Biotransformations: Microbial Degradation of Health-Risk Compounds

Biotransformations: Microbial degradation of health-risk compounds

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edited by M.J. Bull (1st reprint 1983) edited by M.J. Bull edited by M.J. Bull edited by M.E. Bushell Microbial Polysaccharides, edited by M.E. Bushell Modern Applications of Traditional Biotechnologies, edited by M.E Bushell Innovations in Biotechnologie, edited by E.H. Houwink and R.R. van der Meer Statistical Aspects of the Microbiological Analysis of Foods, by B. Jarvis Moulds and Filamentous Fungi in Technical Microbiology, by 0. Fassatiovi Micro-organisms in the Production of Food, edited by M.R. Adams Biotechnology of Animo Acid Production; edited by K. Aida, I. Chibata, K. Nakayama, K. Takinama and H. Yamada Computers in Fermentation Technology, edited by M.E. Bushell Rapid Methods in Food Microbiology, edited by M.R. Adams and C.F.A. Hope Bioactive Metabolites from Microorganisms, edited by M.E. Bushell and U. Grafe Micromycetes in Foodstuffs and Feedstuffs; edited by Z. Jesenski Aspergillus: 50 years on; edited by S.D. Martinelli and J.R. Kinghorn Bioactive Secondary Metabolites of Microorganisms, edited by V. Betina Techniques in Applied Microbiology, edited by B. Sikyta

Biotransformations:

Microbial degradation of health-risk compounds EDITED BY VED PAL SINGH Department of Botany, University of Delhi, Delhi, India

progress in industrial microbiology

ELSEVIER Amsterdam - Lausanne - New York - Oxford - Shannon- Tokyo 1995

ELSEVIER SCIENCE B.V. Sara Burgerhartstraat 25 P.O. Box 21 1, 1000 AE Amsterdam, The Netherlands

L i b r a r y of Congress Cataloging-In-Publicatlon

Data

B i o t r a n s f o r m a t i o n : microbial d e g r a d a t i o n o f health-risk c o m p o u n d s / edited by Ved Pal S l n g h . p. cm. -- ( P r o g r e s s in industrial m i c r o b i o l o g y ; v . 32) I n c l u d e s bibliographical r e f e r e n c e s and index. I S B N 0-444-81977-0 1. X e n o b i o t i c s - - B i o d e g r a d a t i o n . 2. Microbial metabolism. 3. B i o t r a n s f o r m a t i o n (Metabolism.) 4. Xenobiotics--Metabolic I. Singh. Vedpal. 1951- . 11. S e r i e s . detoxicatlon. G!R97.X46B57 1995 628.5'2--dC20 95-1 1660 CIP

ISBN 0-444-81977-0 (Vol. 32) ISBN 0-444-41668-8 (Series) 0 1995 Elsevier Science B.V. All rights reserved

No part of this publication may be reproduced, stored in a retrieval system or transmitted, in any form or by any means, electronic, mechanical, photocopying, recording or otherwise, without the prior written permission of the publisher, Elsevier Science B.V., Copyright & Permissions Department, P.O. Box 521, 1000 AM Amsterdam, The Netherlands. Special regulations for readers in the U.S.A. - This publication has been registered with the Copyright Clearance Center Inc. (CCC), 222 Rosewood Drive, Danvers, MA 01923. Information can be obtained from the CCC about conditions under which photocopies of parts of this publication may be made in the U.S.A. All other copyright questions, including photocopying outside of the USA, should be referred to the copyright owner, Elsevier Science B.V., unless otherwise specified. No responsibility is assumed by the publisher for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions or ideas contained in the material herein. This book is printed on acid-free paper. Printed in The Netherlands

FOREWORD

It is well known that the advancements made in scientific, agricultural, and industrial fields have been responsible for releasing into the environment, waste products, which serve as xenobiotics that pose a potential threat to both h u m a n and animal health. Keeping in view the need for tackling this problem, m a n y thought-provoking ideas from eminent scientists have been compiled in this book, titled "Biotransformations: Microbial Degradation of Health-Risk Compounds", edited by Dr Ved Pal Singh, a scholar of great distinction. Based on his extensive research experience in the field of Applied Microbiology and Biotechnology, Dr Singh has successfully put together, in 14 contributed chapters by recognized experts, a comprehensive and consolidated account of how microorganisms can play a significant role in degrading and detoxifying toxic, carcinogenic, mutagenic, and teratogenic compounds, such as nitrogenous xenobiotics, dimethyl nitrosamine, toxins, haloaromatics, coal-tar, rubbers, tannins, herbicides, pesticides, plastics, polyesters, dyes, and detergents. I strongly feel that the book will have a wide readership, as it will attract academicians, industrialists, professionals, and scientists from various disciplines of diverse interests as well as the students of biology and medicine. Dr Singh's book is the first of its kind, and it will open new vistas of research in the field of Applied Microbiology and Biotechnology in general, and Biotransformations in particular. The book is of very high standard, and I congratulate both Dr Ved Pal Singh and the Elsevier Science Publishers for having brought out such an excellent piece of work through their joint venture.

Professor A.S. Paintal, FRS, FRCP (London), FNA Director General, Indian Council of Medical Research

Former

Delhi, February 6, 1995

DST Centre for Visceral Mechanisms Vallabhbhai Patel Chest Institute University of Delhi Delhi-ll0007, India

vi

About the editor Dr Ved Pal Singh is a Senior Lecturer at the Department of Botany, University of Delhi. He has 13 years of teaching and research experience in the field of Applied Microbiology and Biotechnology, and has about 50 publications to his credit. He is one of the editors for the International Review Series 'Frontiers in Applied Microbiology' and 'Concepts in Applied Microbiology and Biotechnology'. Dr Singh received Young Scientist Awards from the Indian National Science Academy (INSA) and from the United Nations Educational Scientific and Cultural Organization (UNF~CO). He was awarded with the INSA-COSTED Travel Fellowship to visit Hungary (1985). He was a British Council Visitor to the U.K. and Germany (1987). He worked as a Commonwealth Academic Staff fellow at the Royal College, Glasgow (1990, 1991). He chaired sessions and delivered lectures at a number of symposia/seminars/workshops in India and abroad. He has been honoured by the International Society of Conservators and Explorers of Natural Resources (ISCENR), conferring on him the Founder Fellowship with the title FNRS.

This book is dedicated to my teachers Abbas Musavi John Smith and Late Umakant Sinha

o~

VII

PREFACE

In addition to the biological sources of undesirable organochemicals, agricultural and industrial wastes introduce a great variety of xenobiotic compounds in the biosphere and pollute it. Therefore, there is an urgent need to look for the possibilities to tackle this situation of increased agro-industrial wastes generated by fast increasing global population. Microorganisms have tremendous potential to degrade an array of compounds. Owing to their biotechnological potential in degrading and eliminating the hazardous organochemicals, microorganisms occupy a key position in health and environmental protection programmes. "Biotransformations: Microbial Degradation of' Health-Risk Compounds" helps us to u n d e r s t a n d how microbes, following their degradative processes, contribute to the benefit of mankind. It provides a clear u n d e r s t a n d i n g of the biotechnological implications of microbial degradation of health-risk compounds, so as to assist in environmental protection and improve human and animal ihealth. In this book, fourteen chapters contributed by leading scientists from different parts of the world cover a wide variety of xenobiotics such as toxic, carcinogenic, teratogenic, and mutagenic compounds. Moreover, they deal with all aspects of microbial degradation, ranging from screening methods for the degradative microorganisms, processes of degradation, strain improvement for enhanced biodegradation, and elimination of undesirable compounds to improving health and environmental protection strategies. The book intends to provide an opportunity for scientists in the areas of microbiology, biochemistry, engineering, food science, biotechnology, and environmental science to obtain a clear understanding of microbial biotransformations of xenobiotics, and provides an interface between industry and the academic world. I hope that it will provide new dimensions to identify major problems and prospects in Applied Microbiology and Biotechnology, with special reference to Biotransformations and that it will generate new thoughtprovoking ideas for scientists of future generations. I am grateful to the scientists, who accepted my invitation and contributed their valuable review articles for this book. I am greatly indebted to my colleagues Professor N.S. Rangaswamy, Professor K.R. Shivanna, Dr Sudhir Sawhney, Dr A.K. Bhatnagar, Dr S.S. Bhojwani, Dr S.N. Raina, and Dr P.D. Sharma for their valuable suggestions during the course of preparation of the manuscript. I am thankful to Professor A.S. Paintal, Professor Bilquis Musavi, Professor R.P. Roy, and Dr S.K. Chawla for encouragement, and to Dr B.D. Vashishtha, Dr Sarla, and Dr Tripat Kapoor for helping me in various ways. My sincere appreciation is extended to Mr M.S. Sejwal,

o o ~

Vlll

Mr R.I~ Gupta, Mr Krishan Lal, Mr S.I~ Dass, Mr L.I~ Verma, Mr B.I~ Sharma, Mr Ram Pal Giri, and Mr Jai Prakash for technical help. The help rendered by Mr Satish Kumar Sundan and Mrs Mohini Sundan in preparing the manuscript is gratefully acknowledged. I am extremely grateful to the Elsevier Science Publishers B.V., The Netherlands, and especially to Dr Ingrid van de Stadt, the Publishing Editor and her Secretary, Ms Ursula Isaacs for having taken keen interest in my proposal about this book and keeping me well informed about its status from time to time. To complete this book expeditiously, I have received much inducement from my parents Mr Gajadhar Singh and Mrs Anandi Devi, my parentsin-law Mr Biri Singh Bhoj, Mrs Satyawati Devi and Mrs Sushila Devi, and my brothers Mr Prem Pal Singh and Mr Ram Pal Singh. My wife Kusum, daughter Sandhya, and sons Sudhir and Hemant deserve my warmest appreciations for bearing with me throughout the period of my work on this book.

VED PAL SINGH Delhi, 25 January, 1995

ix LIST

AFB 1 AFB 2 AFG 1 AFG 2 AFM 1 AFQ 1 AFR O BOAA CAAT CBAs CBS CF CFCs CIPC CT 2,4-D 1,2-DCA DCAA 1,1-DCE 1,2-DCE DCM DCP DDS DDT DFP DHBA 2,3-DHBPO DMNA 1,3-DNB 2,4-DNP 2,6-DNP 2,4-DNT DPNA DTT FAD FIFRA FMN GCMS GPC GS/GST 4HB ICI

OF

ABBREVIATIONS

Aflatoxin B 1 Aflatoxin B 2 Aflatoxin G 1 Aflatoxin G 2 Aflatoxin M 1 Aflatoxin Q1 Aflatoxin R 0 ~-N-oxalyl L-~,~-diamino propionic acid 2- Chloro- 1 , 3 , 5 - t r i a z i n e - 4 , 6 - d i a m i n e Chlorinated benzoic acids Cyclohexyl benzothiazyl s u l p h e n a m i d e Chloroform Chlorofluoro carbons Isopropyl-N-3-chlorophenyl-carbamate Carbon tetrachloride 2,4-Dichlorophenoxyacetic acid 1,2-Dichloroethane Dichloroacetate 1,1-Dichloroethylene 1,2-Dichloroethylene Dichloromethane Dicumyl peroxide Drug delivery system 1,1,1-Trichloro-2,2'-bis(4-chlorobiphenyl) ethane Diisopropylfluorophosphate Dihydroxybenzoate 2 , 3 - D i h y d r o x y b i p h e n y l dioxygenase Dimethyl nitrosamine 1,3-Dinitrobenzene 2,4-Dinitrophenol 2,6-Dinitrophenol 2,4-Dinitrotoluene N-nitrosodipropylamine Dithiothreitol Flavin adenine dinucleotide Federal Insecticide, Fungicide and Rodenticide Act Flavin mononucleotide Gas chromatographic-mass spectrometry Gel permeation chromatography Glutathione-S-transferase 4-Hydroxybutyrate Imperial Chemical Industries

LAE LAS MCAA MCPA m-DCB MFO MMO MNC MNP NAD NR o-DCB ONP PAGE PAHs p-CB PCBs PCE PCP p-DCB PHA P(3HB) PLFA pMMO PMSF PNP PVC SDS sMMO SMO SOI 2,4,5-T 2,4,6-TNP TCA TCAA TCE TMTD TNT TSCA USEPA VC

Linear alcohol ethoxylate Linear alkylbenzene sulphonate Monochloroacetate 4-Chloro-2-methylphenoxyacetic acid m -D i c h l o ro b e n z e n e Mixed-function oxidase Methane monooxygenase 4-Methyl-5-nitrocatechol m-Nitrophenol Nicotinamide adenine dinucleotide Natural rubber o-Dichlorobenzene o-Nitrophenol Polyacrylamide gel electrophoresis Polycyclic aromatic hydrocarbons p-Chlorobiphenyl Polychlorinated biphenyls Tetrachloroethylene Pentachlorophenol p-Dichlorobenzene P o ly(hy dr oxy alk ano at e) Poly(3-hydroxybutyrate) Phospholipid fatty acid Particulate type of methane monooxygenase Phenylmethylsulphonyl fluoride p-Nitrophenol Polyvinyl chloride Sodium dodecyl sulphate Soluble type of methane monooxygenase Styrene monooxygenase Styrene oxide isomerase 2,4,5-Trichlorophenoxyacetic acid 2,4,6- T r i n i t r o phenol 1,1,1-Trichloroethane Trichloroacetate Trichloroethylene Tetramethyl thiuram disulphide 2,4,6- T r i n i t r ot olue ne Toxic Substances Control Act United States Environmental Protection Agency Vinyl chloride

xi LIST OF CONTRIBUTORS Chapter n,,mbers each contributor

are shown in parentheses

following the

address

of

Todd A. Anderson, Pesticide Toxicology Laboratory, Department Entomology, Iowa State University, Ames, L~, U.S.A. (10)

of

V. A n d r e o n i , Dipartimento di Scienze e Tecnologie Alimentari e Microbiologiche, Universit~ degli Studi di Mihmo, 20133 Milano -Via G. Celoria 2, Italy (1) G. B a g g i , Dipartimento di Scienze e Tecnologie Alimentari e Microbiologiche, Universit~ degli Studi di Mihmo, 20133 Milano -Via G. Celoria 2, Italy (1) S. B e r n a s c o n i , Dipartimento di Chimica Organica e Industriale, Universit~ degli Studi di Milano, 20133 Milan,) - Via G. Celoria 2, Italy (1) Manzoor A. Bhat, Department of Biochendstry and UGC Centre of Advanced Study, Indian Institute of Science,. Bangalore 560012, India (6) J o h n A. Bumpus, Centre for Bioengineering and Pollution Control and Department of Chemistry and Biochemistry, University of Notre Dame, Indiana 46556, U.S.A. (7)

Craig S. Criddle,

National Science Foundation Center for Microbial Ecology, Michigan State University, East Lansing MT 48824, U.S.A. (4)

Yoshiharu Doi, Head, Polymer Chemistry Laboratory, The Institute of Physical and Chemical Research (RIKEN), Hirosawa, Wako-Shi, Saitama 351-01, Japan (9) Jt~rg Fiedler, Universit~it Bielefeld, Facult~it fiir Biologie, Gentechnologie/Mikrobiologie, Postfach 100131, Universit~itsstra~e, D33594 Bielefeld 1, Germany (5)

Karl-Heinz Gartemann, Universit~it Bielefeld, Facult~t fiir Biologie, Gentechnologie/Mikrobiologie, Postfach 33594 Bielefeld 1, Germany (5)

100131,

Universit~itsstra~e,

D-

xii E r w i n Grund, GBF - Gesellschaft fiir Biotechnologische Forschung mBH, Mascheroder Weg 1, D-38124 Braunschweig, Germany (5)

S. Hartmans, Division of Industrial Microbiology, Department of Food Science, Wageningen Agricultural University, P.O. Box 8129, 6700 EV Wageningen, The Netherlands (11,12) Mukesh K. Jain, Department of Civil and Environmental Engineering, Michigan State University, East Lansing MI 48824, U.S.A. (4) S.L. Mehta, Head, Division of Biochemistry, Indian Agricultural Research Institute, New Delhi - 110012, India (13) K a t s u y u k i Mukai, Polymer Chemistry Laboratory, The Institute of Physical and Chemical Research (RIKEN), Hirosawa, Wako-Shi, Saitama 351-01, Japan (9) I.M. Santha, Division of Biochemistry, Indian Agricultural Research Institute, New Delhi - 110012, India (13) R.K. Saxena, Head, Department of Microbiology, University of Delhi, South Campus, New Delhi-ll0021, India (14) A n n e g r e t S c h m i t z , Universit~/t Bielefeld, Fakult~it fiir Biologie, Gentechnologie/Mikrobiologie, Postfach 100131, Universt~tsstra~e, D33594 Bielefeld 1, Germany (5) P. Sharmila, Department of Microbiology, University of Delhi, South Campus, New Delhi-ll0021, India (14) Ved Pal Singh, Department of Botany, University of Delhi, Delhi 110007, India (3,14)

Akio Tsuchii, National Institute of Bioscience and Human-Technology, Agency of Industrial Science and Technology, Tsukuba City, Ibaragi 305, Japan (8) C.S. Vaidyanathan, Department of Biochemistry and UGC Centre of Advanced Study, Indian Institute of Science, Bangalore 560012, India (6) B a r b a r a T. Walton, Environmental Sciences Division, Oak Ridge National Laboratory, Oak Ridge, TN, U.S.A. (10)

xiii David C. White, Center for Environmental Biotechnology, The University of Tennessee, Knoxville, TN, U.S.A. (10) T a d a s h i Yoshinari, Wadsworth Center for Laboratories and Research, New York State Department of Health and School of Public Health, State University of New York at Albany, Empire State Plaza, P.O. Box 509, Albany, NY 12201-0509, U.S.A. (2)

This Page Intentionally Left Blank

xv

CONTENTS Foreword Preface

V

vii

List of abbreviations

ix

List of contributors

xi

1. Microbial degradation of nitrogenous xenobiotics of environmental concern V. Andreoni, G. Baggi and S. Bernasconi 2.

Synthesis and degradation of dimethyl nitrosamine in the natural environment and in humans Tadashi Yoshinari

3. Aflatoxin biotransformations : biodetoxification aspects Ved Pal Singh 4.

Metabolism and cometabolism of halogenated C-1 and C-2 hydrocarbons Mukesh K. Jain and Craig S. Criddle

1

37

51 65

5. Aerobic biodegradation of polycyclic ahd halogenated aromatic compounds Erwin Grund, Annegret Schmitz, Jorg Fiedler and Karl-Heinz Gartemann

103

6. Microbial degradation of halogenated aromatics Manzoor A. Bhat and C.S. Vaidyanathan

125

7. Microbial degradation of azo dyes John A. Bumpus

157

8. Microbial degradation of natural rubber

177

Akio Tsuchii 9.

Microbial degradation of polyesters Katsuyuki Mukai and Yoshiharu Doi

189

xvi

10. Degradation of hazardous organic compounds by rhizosphere microbial communities Todd A. Anderson, David C. White and Barbara T. Walton

205

11. Microbial degradation of styrene S. Hartmans

227

12. Microbial degradation of vinyl chloride S. Hartmans

239

13 . Isolation and characterization of neurotoxin-degrading gene I.M. Santha and S.L. Mehta

249

14. Microbial degradation of tannins R.K Saxena, P. Sharmila and Ved Pal Singh

259

Index

271

Biotransformations: Microbial Degradation of Health Risk Compoututs Ved Pal Singh, editor 9 1995 Elsevier Science B.V. All rights reserved.

Microbial degradation of environmental concern

nitrogenous

xenobiotics

of

V. Andreoni a, G. BaggP and S. Bernasconi b aDipartimento di Scienze e Tecnologie Alimentari e Microbiologiche, Universit~ degli Studi di Milano, 20133 Milano - Via G. Celoria 2, Italy bDipartimento di Chimica Organica e Industriale, Universit~ degli Studi di Milano, 20133 Milano - Via G. Celoria 2, Italy

INTRODUCTION Nitrogen forms a variety of functional groups in combination with carbon, hydrogen, and oxygen. These functional groups have been particularly useful for adapting and activating aromatic compounds to be used as chemical intermediates in synthetic processes [1]. In addition, many final products, such as pesticides, explosives, drugs, dyes, antioxidants and antiozonants, contain nitrogen functionalities [2-7]. Consequently, these compounds are contaminants of rivers, ground water, soils treated with pesticides, and atmosphere [8-11]. Exposure to amines and related compounds has large impact on human health: workers exposed to benzidine and naphthylamlne have developed cancer of the bladder [12,13]; the Food and Drug Administration (F.D.A.) has found that aromatic amines [14] and nitroaromatics [15] can enter the food chain. In addition, many degradation products from nitroaromatics are easily polymerized, in presence of oxygen, to persistent macromolecules [16]. The wide distribution of these compounds in the environment, coupled with their toxicity, has given rise to concern about their environmental fate. The complete degradation of nitrogenous compounds is mainly the result of microbial attack and represents one of the primary mechanisms by which these pollutants are eliminated from the environment. The term d e g r a d a t i o n is olden used only to indicate the disappearance of a compound, which, in turn, is transformed into another, with no evidence of the extent of degradation. The degree of degradation depends on the nature of the compounds: some are resistant to microbial attack, others are partially broken down into persistent intermediates, or transformed into more toxic products. On the contrary, complete biodegradation will result in mineralization to carbon dioxide or methane with release of nitrite or ammonium ion. In aerobic environment, 02 is both the terminal electron acceptor and a reactant in the initial reactions. In the absence of oxygen, however, organic compounds, like nitrate, sulphate and carbonate, are alternate electron acceptors in the microbial degradation of organic material, and

the presence or absence of these electron acceptors plays a crucial role in biodegradability and influences microbial activity and diversity. The intent of this review is to present a broad and updated overview of the physiological, biochemical, and genetic basis of biodegradation of nitrogenous compounds by aerobic and anaerobic microorganisms.

NITROAROMATIC S Nitroaromatic compounds are produced industrially on a large scale. Such chemicals are widely used as pesticides, or for other chemical uses [2-7]. Nitroaromatics m a y be produced enzymatically in microbial cultures [17], or photochemically in urban air [18,19]. Nitroaromatics are highly toxic to m a n and mammals, being easily reduced by enzymes to nitroso and hydroxylamine derivatives. These derivatives m a y lead to the formation of either metahemoglobin, which is unable to bind oxygen, or of nitrosoamines, which are carcinogenic [14,20,21]. Some nitroaromatics, such as nitropyrene [19], are mutagenic and several nitrophenols have an uncoupling effect on oxidative phosphorylation [22]. Most nitroaromatics are highly toxic also to bacteria and, consequently, m a y inhibit microbial growth. In activated sludges, their presence m a y destabilize the continuous process of sewage treatments. Nitroaromatics are slowly degraded by microorganisms both under aerobic and anaerobic conditions, and the metabolic steps involved in the degradation have been poorly docllmented until now. Two major catabolic pathways are involved in the degradation of nitroaromatics (Figure 1) [23]. In the first pathway, the nitro group is reduced to an aniline intermediate, which is further degraded to a m m o n i u m ion and catechol [5,24-28]. A reduction of nitro substituent, under both aerobic and anaerobic conditions, seems to be a c o m m o n enzymatic mechanism in the environment [5,29]. Such reduction has been demonstrated in various organisms which utilize the nitro compound as an electron acceptor. The activity of nitroreductases, m a n y of which have a broad substrate specificity, has been demonstrated in cell-flee systems, and some enzymes have been purified and characterized [5,19,29]. The resulting aromatic amines are often further transformed into persistent azo compounds or polymers by biotic or abiotic processes [1,30,31]. In the second pathway, the nitro substituent is directly removed as nitrite [24,32,33], with the formation of catechol. The microbial degradation of nitroaromatics to catechol involves a series of reductions and oxidations, generally catalyzed by reductases and oxygenases.

O2,2[H]" NH4+_--.... ~ ~ %'~'~

X

~2.

|

X

OH

X

2[HI

Ring cleavage --,----------,-and mineralization

/ o

,'

Figure 1. Microbial degradation of nitroaromatic compounds [23].---, steps demonstrated;__, steps postulated; A, nitroreductase; B, aniline oxygenase; C, nitrophenyl oxygenase; D, chinoreductase? Nitrotoluenes 2,4,6-Trinitrotoluene (TNT) is the predominant conventional explosive used by military forces [34], and the disposal of wastes containing TNT leads to soft, sediment, and water contamination [35]. This is of great concern because TNT causes liver injury and marked changes in the hemopoietic system, producing anemia in humans and other m~mmals [36]. Moreover, TNT is toxic to certain fish at concentrations greater t h a n 2 ~tg/ml [37] and to certain green algae [38]; finally, TNT is mutagenic [5]. TNT was shown to be oxidized by three Pseudomonas-like bacteria; the degradation was accelerated by the addition of glucose or yeast extract and proceeded through the formation of several intermediates: dinitrohydroxylaminotoluene, dinitroamino toluene, nitrodiaminotoluene, and azoxy toluenes [39]. Among these metabolites, only nitrodiaminotoluene, and dinitroaminotoluene were not degraded further. The same reduced and azoxy compounds have been isolated by McCormick et al. [5] (Figure 2). They found that the nitro groups of TNT were reduced by both aerobic and anaerobic systems, and that the number of the nitro groups reduced depended on the reducing potential of the system and on the species utilized. Cell-free extracts of Veillonella alcalescens utilized 3 moles of H 2 t o reduce 1 mole of nitro group. The

CH3 O2NQNO2

O2Nj~NO2 ~',,~,~CH3 N

O~N

II

II

CH3 O2NQ N O 2

~

CH3 02 N~"~,,~NO2

Vlll

N N -.--,-- O

NO2

"~

OzN~

CH3 N HOH V

II

CH3 J O 2 N Q NOz

NO2

O2N~O 2 CH3 VII

NHOH

1

CH3

O2NQNH2 NO2

III O2NO

Vl

-,,,.

IV J CH3

CH3

NO2

NH2

O2NQNH2 NH2

1 CH3 H2NQNH2

NH 2 IX Figure 2. Proposed pathway for transformation of 2,4,6-TNT [5]. I, 2,4,6TNT; II, 4-hydroxylamino-2,6-dinitrotoluene; III, 4-amino-2,6dinitrotoluene; IV, 2,4-diamino-6-nitrotoluene; V, 2-hydroxylamino-4,6dinitrotoluene; VI, 2-amino-4,6-dinitrotoluene; VII, 4,4'-azoxycompound; VIII, 2,2'-azoxycompound; IX, 2,4,6-triaminotoluene. oxidation of the hydroxylamino derivatives to azoxy compounds may also occur non-enzymatically in anaerobic environment [40]. The same reduced products are present in the urine excreted by rabbits, rats, or human volunteers fed with TNT [41]. TNT is also degraded by different fungi [42,43]. Rhizopus stolonifer was able to degrade almost all TNT, when present in cultural broth at a concentration of 100 rag/1 [42], and extensive biodegradation of ["C]-TNT by the white rot fungus Phanerochaete chrysosporium was also observed [44]. Biodegradation of [14C]-TNT occurs even in a mixture of soil and corncobs

inoculated with P. chrysosporium [44]. However, s u b s t a n t i a l l y less [14C]-TNT was converted to 14CO2 in soil cultures, compared with liquid cultures. 2,4-Dinitrotoluene (2,4-DNT), listed as a priority pollutant by the United States Environmental Protection Agency (USEPA) [45], is the major impurity resulting from the manufacture of TNT, and is a starting material for the synthesis of toluenediisocyanate, used in the production of polyurethane foam. 2,4-DNT was transformed to 2-amino-4-nitrotoluene, 4-amino-2-nitrotoluene, 2-nitroso-4-nitrotoluene and 4-nitroso-2nitrotoluene by a mixed culture derived from activated sludge only under anaerobic conditions and with an exogenous carbon source [46]. The two nitroso compounds were unstable and could be detected between 48 and 72 h of incubation. A Pseudomonas sp., which is able to degrade aerobically 2,4-DNT, using the latter as the sole source of carbon and energy with stoichiometric release of nitrite, has been described. 4-Methyl-5-nitrocatechol (MNC) accumulated transiently when cells grown on acetate were transferred to medium containing 2,4-DNT. Conversion of 2,4-DNT to MNC was catalyzed by a dioxygenase [47] (Figure 3). MNC was then rapidly oxidized with the removal of the second nitro group as nitrite.

F NO2

CH3 1 H

L

-

NO~ HO

NO2

.._1

OH

Figure 3. Initial steps in 2,4-DNT degradation pathway [47]. Finally, nitrotoluenes largely used in the manufacture of azo and sulphur dyes, and in the production of explosives [48], have been detected at high levels in waste waters from paper mills and chemical plants [45]. Delgado et al. [49] showed that the biotransformation of nitrotoluenes into more oxidized nitroaromatic chemicals is mediated by the upper pathway of the TOL-plasmid. In fact, they found that the TOL-upper-pathways enzymes recognize nitroaromatics as substrates, although the regulator, the XyLR protein, does not recognize nitrotoluenes as effectors. The TOL-encoded toluene monooxygenase enzyme biotransformed 3-nitrotoluene and 4-nitrotoluene into their corresponding

benzyl alcohols and b e n z a l d e h y d e s , but not 2-nitrotoluene. The t r a n s f o r m a t i o n of nitrobenzyl alcohol into the corresponding nitro benzaldheyde was carried out by the same toluene monooxygenase, in agreement with Harayama et al. [50], who have reported that this enzyme, in addition to its primary oxidative activity, also shows an alcohol dehydrogenase activity. Recently, it has been reported that cells of Pseudomonas putida F1 and Pseudomonas sp. strain JS150 were capable of degrading nitrotoluenes into 3-methyl-6-nitrocatechol and 2-methyl-5-nitrophenol through initial oxidation into 4-nitrotoluene2,3-dihydrodiol by toluene dioxygenase [51].

Nitrobenzenes Nitrobenzenes are widely used in the manufacture of aniline and pyroxylin compounds, in the refinery of lubricant oils and in the production of soap and shoe polishes [52]. 1,3-Dinitrobenzene (1,3-DNB) is the main impurity of TNT [53]. These non polar nitroaromatic compounds are considered recalcitrant to microbial attack [54] for their resistance to the reduction of electron density in the aromatic ring, which can hinder electrophilic attack by oxygenase, and for their toxicity against microorganisms [4,5]. However, several bacterial strains, capable of degrading toluene and to oxidize nitrobenzene, have been isolated. While in cells of Pseudomonas putida F1 and Pseudomonas sp. strain JS150, a dioxygenase mechanism converts nitrobenzene into the corresponding dihydrodiol, in other microorganisms, a monooxygenase is instead responsible for the initial attack on nitrobenzene [55]. Recently, a new Rhodococcus species, isolated under nitrogen limiting conditions from contomlnated soils, and capable of utilizing 1,3-DNB, has been described. 0.5 mM of 1,3-DNB was completely and immediately metabolized by induced cells with release of 2 moles of nitrite per mole of 1,3-DNB via 4-nitrocatechol [56] (Figure 4). According to the mechanism

NO2

~~]~NO2

NO2

NO 2 .OH H OH

NO 2

NO2

OH

-- ~-~TCC

OH

Figure 4. Proposed pathway for degradation of 1,3-DNB by sp. QT-1 [56].

NO2

Rhodococcus

reported in Figure 4, 4-nitrocatechol could be generated by an initial 3,4-dioxygenation with subsequent elimination of 1 mole of nitrite. Interestingly, Rhodococcus utilized 1,3-DNB as source of nitrogen in the absence as well as in the presence of high amounts of ammonium ion.

Nitrophenols Nitrophenols, used in the manufacture of dyes, explosive, and pesticides [2,5,6], are released into the environment during the hydrolysis of several organophosphorous pesticides, such as parathion. 2-Nitrophenol, 4-nitrophenol, and 2,4-dinitrophenol are priority pollutants according to the USEPA [57]. 2,4-Dinitrophenol (2,4-DNP) is an uncoupler of electron transport [22,58,59] and its structural analogues, 4,6-dinitro-2-methylphenvl, 2sec-butyl-4,6-dinitrophenol and Dinoseb are important pesticides [60]. Dinoseb, which is also the major degradation product of the herbicide, Acrex by soil microorganisms [61], is responsible for h e a l t h and e n v i r o n m e n t a l hazards [62]. p-Nitrophenol (PNP) was degraded either by resting or growing cells of a Flavobacterium strain with stoichiometric release of nitrite and formation of 4-nitrocatechol [63]. The degradation of PNP was accompanied by the disappearance of the characteristic yellow colour in the medium, indicating cons!lmption of the nitrogen, when nitrophenols were tested as nitrogen source. The cells grown on PNP also oxidized m-nitrophenol into nitrohydroquinone, but did not use m-nitrophenol as carbon source for growth. Rapid biodegradation of PNP was shown to occur in a pond in 6 days; a second treatment of the pond with PNP enhanced its biodegradation which began immediately [64]. Differently, a Moraxella strain degraded PNP by replacing the nitrogroup with a hydroxyl group and accumulating traces of hydroquinone in the medium. Hydroquinone was then converted into ~-ketoadipic acid via T-hydroxymuconic semialdehyde [65]. A Pseudomonas putida utilized o-nitrophenol (ONP) and m-nitrophenol (MNP) as source of carbon and nitrogen, but not PNP. Growing cells of these organisms degraded ONP and MNP, releasing nitrite and ammonium, respectively. The enzymes involved in the metabolism of ONP or MNP were inducible. Only the degradation pathway of ONP has been described by using a crude enzyme extract. The crude extract converted ONP to nitrite and catechol, which were further metabolized through the orthocleavage [32]. Zeyer et al. subsequently observed that P. putida did not utilize parasubstituted derivatives of ONP and, in the same study, they characterized the inducible nitrophenol oxygenase responsible for the release of nitrite [66]. The enzyme was found to be soluble, NADPH-dependent

and its activity was stimulated by magnesium and manganese ions, but not by FAD, and consisted of a single polypeptide chain with a molecular weight of 58,000 (determined by gel filtration) or 65,000 (determined on a sodium dodecyl sulphate polyacrylamide gel) [67]. The degradation of PNP was found to occur also under cometabolic conditions by a Pseudomonas sp. in presence of glucose, but not in presence of phenol that, instead, inhibited PNP mineralization [68]. The oxidative elimination of nitrite ions by dinitrophenols has been detected also during the mineralization of 2,6-dinitrophenol (2,6-DNP) by Alcaligenes eutrophus JMP 134 [69]; a total degradation of 2,6-DNP was performed by a Pseudomonas strain, which, although grew scant, decolourized the culture medium within 3 to 4 days of incubation [70]. Under nitrogen-limiting conditions, two Rhodococcus erythropolis strains, able of mineralizing 2,4-DNP as the sole source of carbon, have been isolated. Both strains metabolized 2,4-DNP, present at a final concentration lower t h a n 0.5 mM with liberation of stoichiometric amounts of nitrite and of low amount of 4,4-dinitrohexanoate [71]. The identification of the last compound as the only organic metabolite of 2,4-DNP suggested the involvement of a reductive mechanism in the degradation pathway. The initial reduction of the aromatic ring gives evidence for a nucleophilic attack in consequence of the highly electrophilic character of the aromatic nucleus of 2,4-DNP, that favours an initial reductive reaction. A different reductive mechanism was described for the degradation of 2,4-DNP by a Fusarium oxysporum strain [72]. Although 2,4-DNP is used extensively in fungicidal preparations, Fusarium reduced it to the less toxic 2-amino-4-nitrophenol and its isomer 4-amino-2-nitrophenol. The reduction of the nitro groups occur in successive stages by the intermediate formation of the nitroso and hydroxyleamino groups. Finally 2,4,6-trinitrophenol (2,4,6-TNP), picric acid, is used as explosive under the form of ammonium 2,4,6-trinitrophenoxide. Both picric acid a n d picramic acid (2-amino-4,6-dinitrophenol), deriving from the bioconversion of picric acid under anaerobic conditions, are well-known mutagenic compounds [73]. The degradation of picric acid proceeded by nucleophilic attack of the aromatic ring [74]. A spontaneous mutant of Rhodococcus erythropolis HL 24-2, originally isolated for its ability to degrade 2,4-dinitrophenol [14], could also utilize picric acid as nitrogen source. The mutant HLPM-1 transiently accumulated an orange-red metabolite, which was identified as a hydride-Meisenheimer complex of picric acid. This complex was further converted with release of nitrite. 2,4,6- Trinitrocyclohexanone was the dead-end metabolite of the degradation of picric acid.

Nitro and aminobenzoates Like all nitroaromatics, these compounds e n t e r i n d u s t r i a l waste streams, and may accumulate in the environment. In aerobic metabolism of 2- and 4-nitrobenzoate, reduction of the nitro group via nitroso and hydroxylaminobenzoate was d e m o n s t r a t e d [75-78]. In this case, the nitro group was used as a terminal electron acceptor. The amino intermediate was transiently accumulated during the growth on the corresponding nitrobenzoate, but it was not clear whether or not amines were intermediates in aerobic degradation of nitrobenzoates. Ke et a]. [79], working with a strain of Flavobacterium capable of mineralizing 2-nitrobenzoate, indicated t h a t the cells growing on 2-nitrobenzoate were not s i m u l t a n e o u s l y a d a p t e d to 2 - a m i n o b e n z o a t e . A similar consideration was made in the case of the degradation of 4-nitrobenzoate [76]; 4-aminobenzoate did not lie on the direct oxidative p a t h w a y because no appreciable oxidation of this substrate was observed with cells grown on 4-nitrobenzoate. Involvement of amines, however, is often implied. The incubation of cells of Pseudomonas sp. CBS3 with the isomers of nitrobenzoate, under aerobic conditions, allowed to detect the presence of the corresponding amines in cultural broths. In the course of the conversion of 4-nitrobenzoic acid, 4-nitrosobenzoic acid appeared shortly after the start of the reaction, and was no longer detectable after 24 h of incubation, when the major product present was instead 4-aminobenzoic acid. As the same conversion of 4-nitrobenzoate by Pseudomonas sp. CBS3 occurred in absence of molecular oxygen, the nitroaromatic compound may be electron acceptor, according to the scheme proposed in Figure 5 [80]. Recently, t h e d e g r a d a t i o n of 4 - n i t r o b e n z o a t e by a Comamonas acidovorans NBA-10 was shown to proceed via reduction of the nitro group, but without the formation of 4-aminobenzoate. Cell extracts, in presence of NADPH, degraded 4-nitrobenzoate into 4-hydroxylamino-

NO 2

NO

NHOH

NH 2

Figure 5. Proposed pathway for the reduction of nitroaromatic by Pseudomonas sp. CBS3: R, -OH, -COOH, -C1, -NO 2 [80].

compounds

10 benzoate and 3,4-dihydroxybenzoate [81] (Figure 6), evidencing a new pathway for its aerobic degradation which did not involve molecular oxygen [82]. The conversion of 2-nitrobenzoate by an oxygenative reaction, yielding nitrite, is reported by Andreoni et al. [83]. An Achromobacter strain utilized 2-nitrobenzoate as only carbon and nitrogen source with release of nitrite in the culture medium. Simultaneous oxidation of 2nitrobenzoate and 2,3-dihydroxybenzoate by resting cells of Achromobacter suggests the involvement of an inducible dioxygenase in the degradation of 2-nitrobenzoate. 2-Aminobenzoic ( a n t h r a n i l i c ) acid is an i m p o r t a n t i n t e r m e d i a r y metabolite in both biosynthetic and catabolic pathways of microorganisms. In fact, it serves as precursor for tryptophane for many bacteria and molds [84,85], and is formed by bacterial reduction of 2-nitrobenzoic acid [76,77], and in the degradation of compounds containing an indole moiety, both aerobically [86, 87] and anaerobically [88]. While 2- and 4a m i n o b e n z o a t e s occur in n a t u r e , 3 - a m i n o b e n z o a t e is a xenobiotic compound, produced mainly for the synthesis of azo dyes. A toxic effect of 3-aminobenzoate has been reported for man and many mammals, which metabolize it partly to 3-ureidobenzoic acid and 3-aminohippuric acid [89]. The aerobic catabolism of Aminobenzoates proceeds via catechol [90] or gentisic acid [91], and requires molecular oxygen. Brown and Gibson reported some denitrif~ng organisms belonging to the genus Pseudomonas, t h a t degraded 2-aminobenzoate completely into CO 2 and NH4+ [92]. Nitrate, the terminal electron acceptor, was first reduced to nitrite and then to nitrogen. Aerobically, 3-aminobenzoate was degraded through hydroxylation to 5-aminosalicylic acid which was further metabolized (Russ, Stoltz and Knackmuss, Abstr. Ann. Meetg. VAAM, Freiburg, 1991; Poster P 136). Anaerobically, 3-aminobenzoate was found to be decomposed both by a sulphate-reducing bacterium in pure culture, and by a methanogenic and a nitrate-reducing enrichment culture [93]. The

COOH

NO2

NADPH L~NADP''-~

COOH

NO

NADP N~ADP+~--

COOH

HNOH

COOH ~N H3~

OH

OH

Figure 6. Proposed degradative pathway of 4-NBA by Comamonas acidovorans NBA-IO [81].

11 sulphate reducer oxidized 3-aminobenzoate completely to CO 2, with concomitant reduction of sulphate to sulphide and release of ammonium. In the absence of an external electron acceptor, 3-~minobenzoate was degraded by a methanogenic culture, consisting of three types of bacteria into CO 2, CH4, and NH4§ The consortium was constituted by a short rod able to ferment 3-aminobenzoate, a H2-degrading Methanospirillumlike bacterium, and an acetate-degrading Methanothrix-like bacterium.

NITRILES

Nitriles are cyanide-substituted carboxylic acids, which occur naturally and synthetically, and are of the general structure, R-CN. The naturally occurring nitriles are found in higher plants [94-97], bone oils, insects [98], and microorganisms [99,100]; the synthetic ones are used industrially in benzonitrile herbicides [101], as organic solvents and in the synthesis of polymers, plastics [102,103], synthetic fibers, resins, and dye stuffs. There is little information on the ecological impact of these compounds, most of which are highly toxic, and some mutagenic and carcinogenic. Particularly, acrylonitrile is a largely used vinyl cyanide of which the production in U.S. was estimated at 2.5 x 109 pounds [104-106]. Because of its acute neurotoxicity, mutagenicity, and teratogenicity, the USEPA has targeted acrylonitrile as a priority pollutant [56]. Microbial degradation of nitrile compounds was performed by different microorganisms, capable of growing on various aliphatic and aromatic nitriles [107-109]. They are degraded through two pathways (Figure 7): one is the direct hydrolysis of nitriles to carboxylic acid and ~mmonia, catalyzed by nitrilases. Nitrilases, that utilize benzonitrile and related aromatic nitriles as substrates, have been purified from Pseudomonas sp. [110,111], Nocardia sp. strains NCIB 11215 [112] and NCIB 11216

R1CN

nitrilase 2 ~ H20 ~ R1COOH+NH3

nitrile amidase R2CN hydratase ~ R2CONH2

So

R2COOH+ NH3

Figure 7. Proposed pathway for the degradation of nitriles: R1, -phenyl; -~-~-alkenyl; R2, -alkyl.

12 [113], Fusarium solani [114], Arthrobacter sp. [115], Escherichia coli, transformed with a Klebsiella ozaenae plasmid DNA [116], Rhodococcus rhodochrous J1 [117,118], and Alcaligenes faecalis JM3 [119, 120]. The enzyme of Rhodococcus rhodochrous J1 was employed for the production of p-aminobenzoic acid from p-aminobenzonitrile [121], and nicotinic acid from 3-cyanopyridine [122]. The conversion of 3-cyanopyridine to nicotinic acid, by a nitrilase of Nocardia rhodochrous LL100-2, was also reported by Vaughan et al. [123]. These nitrilases were usually inactive on aliphatic nitriles. More recently, a new nitrilase, that acts preferentially on aliphatic nitriles, was purified and characterized in Rhodococcus rhodochrous K22 [124]. The other pathway, working preferentially on aliphatic nitriles, is a two-step degradation process, involving nitrile hydratase and amidase, via an amide as intermediate. The corresponding amides are then hydrolyzed into the respective carboxylic acids and ammonia [108,125127]. Aliphatic nitrile hydratases, that catalyzed the hydratation of nitriles to amides, were purified and characterized in Arthrobacter sp. J1 [128], Brevibacterium R312 [129], and Rhodococcus sp. N774 [130]. In the first strain, the activity of an amidase, which forms acetic acid and ammonia stoichiometrically from acetamide, was also detected [131]. Bioconversion of dinitrile to mononitrile catalyzed by nitrile hydratase and amidase was obtained from Corynebacterium sp. C5. The two enzymes were constitutively formed in cells [132]. The production of amides from nitriles has been studied by several workers, and most of them focused on the accumulation of acetamide from acetonitrile [126,133-136]. The enzymatic production of acrylamide from acrylonitrile by nitrile hydratase of P. chlororaphis B23, Rhodococcus sp. N-774, and Klebsiella pneumoniae, respectively has been reported [137-142]. These microorganisms exhibited a high nitrile h y d r a t a s e activity and a low amidase activity, allowing the accumulation of the corresponding amide. Nagasawa et al. optimized the reaction conditions for the production of nicotinamide by a nitrile hydratase, found in Rhodococcus rhodochrous J1. The enzyme contains cobalt, and shows high activity towards 3-cyanopyridine [143,144]. Recently, the potential of bacterial enzymes for the synthesis of aromatic, optically active amides, and carboxylic acids from racemic nitriles was evaluated. An enantiomer-selective amidase, active on several 2-aryl and 2-aryloxy propionamides, was identifided and purified from Brevibacterium sp. strain R312 [145]. A nitrilase, found in Acinetobacter sp. strain AK226 and able to hydrolyze efficiently both aromatic and aliphatic nitriles, was reported to hydrolyze racemic nitriles to optically active 2-aryl propionic acids [146]. Enzyme system of Rhodococcus butanica could be successfully adapted for the kinetic resolution of a-arylpropionitriles resulting in the formation of (R)-

13 amides and (S)-carboxylic acids [147]. Finally, from racemic mandelonitrile and its acetylated derivatives, R(-)-mandelic and R(-) acetylmandelic acids were obtained by using enantioselective nitrilases of Alcaligenes faecaZis ATCC 8750, and Pseudomonas, respectively [148,149]. The interest of some workers has been focused on the biodegradative process of halogenated aromatic nitriles, herbicides, widely used in post-emergence control of seedling broadleef weeds in a number of t o l e r a n t p l a n t s [150]. The di-ortho-substituted Dichlobenil (2,6dichlorobenzonitrile) is a persistent compound [151], presumably because the double ortho-substitution in the aromatic ring is incompatible with enzyme attack. To confirm this hypothesis, Dichlobenil was not attacked by nitrilase either of Nocardia [113] or of Fusarium solani [114], both found capable of growing on benzonitrile as sole carbon and nitrogen source. On the other hand, the same fungal enzyme was tested on different benzonitriles such as Yoxynil (3,5-diiodo-4-hydroxybenzonitrile) and Bromoxynil (3,5-dibromo-4-hydroxybenzonitrile) [114], hydrolyzed at significant rates. The metabolism of Bromoxynil has been studied in some microorganisms, revealing the presence of amide and acid products [3,152-154] (Figure 8). KlebsielZa pneumoniae subsp, ozaenae was shown to transform completely the herbicide, with the involvement of a nitrilase enzyme to

CONH2

f

CN 1

OH Bromoxynil

~~L~Br OH 3,5-dibromo-/4- " ~ hydroxybenzamide

COOH Br

OH 3,5-dibromo-4hydroxybenzoicacid

Figure 8. Initial reactions in the metabolism of the herbicide, Bromoxynil

[3].

14 corresponding acid, and to utilize the liberated ammonia as sole nitrogen source. As the cyano-moiety of the molecule is important for the toxic properties of the herbicide, its removal is essential for the detoxification of the compound [3]. HETEROCYCLIC

COMPOUNDS

Indole Indole and its derivatives form a class of toxic recalcitrant compounds released into the environment through cigarette smoke, coal-tar, and sewage. Many of these compounds, which are present in several edible plants, are responsible for different diseases in cattle and goats [155, 156], and have shown to be toxic and mutagenic [157,158]. The studies on the microbial metabolism of indole always indicate anthranilic acid as intermediate, but the degradative pathways observed, leading to this compound, were found to be different. By carrying out oxygen uptake studies, Sakamoto et al. proposed indoxyl, dihydroxyindole, and isatin as metabolites during the bacterial degradation of indole [159], whereas another group suggested t h a t indole is metabolized via 2,3-dihydroxyindole, through the initial formation of an epoxide. Dihydroxyindole is converted into anthranilic acid by dihydroxyindole oxygenase, an inducible enzyme, which appears only when the organism has grown on indole. This conversion is apparently a single enzymatic step [160]. More recently, Ensley et al., by using cloned naphthalene dioxygenase and other dioxygenases, reported that indole oxidation is a property of bacterial dioxygenases, that form cisdihydrodiols from other aromatic hydrocarbons by a dioxygenation reaction [161]. Spontaneous elimination of water from the cis-dihydrodiol would yield indoxyl, the precursor of indigo. Indoxyl was found as an intermediate also in the metabolism of indole by Alcaligenes. On the basis of its respiratory activity, isatin is considered the next intermediate, while in broth cultures with indole, anthranilic acid was found. This latter intermediate was catabolized via gentisate, which is cleaved by g e n t i s a t e 1,2-dioxygenase, p r e s e n t in cells grown on indole, to maleylpyruvate [162]. A strain of AspergiUus niger, which cometabolized indole in the presence of glucose and nitrate, monohydroxylated this compound into indoxyl (Figure 9). This mechanism is considered prevalent in fungi and other higher organisms as well as in anaerobic bacteria, which hydroxylate indole to oxindole, whereas in aerobic bacteria unhydroxylated aromatic compounds are attacked by dioxygenases [161]. Indoxyl was further converted to N-formylanthranilate by a dioxygenase, but this activity was not demonstrated for the instability of the substrate. In the cytosolic fraction, N-formylanthranilate deformylase, anthranilate hydroxylase, dihydroxybenzoate (DHBA) decarboxylase, and catechol

15

H Indole

H Indoxyl

H N-formyl anthranilate

Indigo ortho ~ ring

cleavage

~

(~ , OH

OH

OH

1 COOH OH

~COOH v

-NH2

Figure 9. Proposed pathway for t h e degradation of indole by A. niger. Indoxyl is a proposed intermediate [87]. dioxygenase, induced by growth on the glucose plus indole, were detected. DHBA decarboxylase has been found only in fungal systems. The anaerobic metabolism of indole and its derivatives has been studied more recently, and relatively little is known about the fate of these compounds in anaerobic conditions. Wang et al., using methanogenic consortia, reported biodegradation of indole into methane and carbon dioxide u n d e r strict anaerobic conditions [163], and oxindole was recognized as the initial intermediate [164,165] (Figure 10). The increase in net m e t h a n e production indicates t h a t indole was mineralized presumably through the formation of anthranilic acid. The ability of sediment and sewage sludge microcosmos to degrade indole was dependent upon several factors, including incubation temperature and the amount of sediment or sludge inoculum used [165]. The effect of substituent groups on indole hydroxylation reactions was studied by Gu and Berry with an indole-degrading methanogenic consorti,!m, that was also able to d e g r a d e 3 - m e t h y l i n d o l e and 3-indo|yl a c e t a t e [166]. Oxindo|e, 3-methyloxindole, and indoxyl were identified as metabolites of indole

16 H I/H N H indote

~--

I~N

I~o H H oxindole tautomer

CH/~+ CO2

~--'---~ ~ N . ~ O H oxindole

~

COOH NH 2

Figure 10. Suggested pathway of methanogenic indole fermentation [164]. and its two derivatives, respectively. Isatin was produced as intermediate when the consortium was amended with oxindole, providing evidence that degradation of indole proceeded through successive hydroxylations of C-2 and C-3 atoms prior to ring cleavage. The 3-substituted indoles were not further metabolized by the consortium. The metabolism of indole was also explored with a denitrifying microbial community. After oxindole and isatin formation, the addition of two hydroxyl groups has been postulated to yield isatoic acid, which is chemically unstable and spontaneously decarboxylates to form anthranilic acid. Dioxindole was also isolated from the denitrifying cultures exposed to isatin, presumably by reducing agents present in the sewage-sludge inoculum; however, whether dioxindole was formed by chemical or by microbiological process remains uncertain, and it has not been clarified if the dioxindole formed is reoxidized to isatin or if the reduction step represents a branch point in the metabolism of indole [167]. The degradation of indole, under sulphate-reducing conditions, has not been elucidated todate; only Bak and Widdle have isolated, from a marine enrichment with indole as sole electron donor and carbon source, a sulphate-reducing bacterium, ascribed as new species of Desulfobacterium indolicum, which oxidized completely indole to CO 2 with sulphate as electron accepter [168]. S-Triazines Among heterocyclic compounds, S-triazines are of great environmental concern, as different compounds, largely used, derive from their nucleus. All S-triazines, used as herbicides, are diamino-S-triazines bearing a chloro, methoxy or methylthio group, linked directly to the nucleus. Among these compounds, atrazine (2-chloro-4-ethylamino-6-isopropyl-

17 amino-l,3,5-triazine) has often been detected in groundwater and soil because of its persistence [169]. Though physico-chemical decomposition plays an important role in the removal of these compounds from the environment [170], microbial degradation is the principal mechanism of

detoxification. The metabolism of chloro-S-triazines in soil involves reactions of dealkylation, deamination, hydroxylation, and ring cleavage [171]. Dealkylation of chloro-S-triazines does not remove their toxicity, which has been, instead, attributed to the release by chemical hydrolysis of active chlorine [172]. In a study on the effects of atrazine and its degradation products on phototrophic microorganisms, the most toxic degradation product was deethylated atrazine, which was 2 to 7 times more effective towards cyanobacteria than deisopropylated atrazine. On the contrary, diamino and hydroxyatrazine were non-toxic [173]. The first demonstration of the microbial degradation of chloro-Striazines is the fungal dealkylation of simazine (2-chloro-4,6-blsethylamino-l,3,5-triazine) to deethylsimazine, and to another product, tentatively identified as 2-chloro-l,3,5-triazine-4,6-diamine (CAAT) [174,175]. Analogously, the same fungus, Aspergillus fumigatus Fres. monodealkylates atrazine to 2-chloro-4-amino 6-isopropylamino-Striazine and 2-chloro-4-ethylamino-6-amino-S-triazine [176] (Figure 11). Cook and Hiitter reported that deethylsimazine was quantitatively utilized as a nitrogen source by a strain of Rhodococcus corallinus, yielding ethylamino-dihydroxy-triazine which is utilized in co-culture with a Pseudomonas to yield cell material [177]. The same authors confirmed that the initial reaction is a quantitative hydrolytic ring dechlorination by two isofunctional, but different, hydrolases [178]. CAAT, the other product of fungal dealkylation of simazine, was degraded to 2-chloro-4-amino-l,3,5-triazine-6(5H)-one, a new product in the biodegradation of chlorinated S-triazines, which appeared to be further metabolized [179]. In this mechanism, the initial dechlorination, observed with deethylsimazine, does not occur [177]. Giardina and coworkers [4,180,181], working with a Nocardia strain capable of growing on atrazine, isolated dealkylated and deaminated products of the herbicide, among which 4-amino-2-chloro-l,3-5-triazine represented the most highly degraded compound, still having chlorine in the triazinic ring. This metabolite does not accumulate in the medium, as it undergoes rapid hydrolysis, causing the cleavage of the heterocyclic ring. These results suggest that the microbial attack is the necessary step for further chemical transformations, leading to ring cleavage. Behki and Khan isolated Pseudomonas species, that caused N-dealkylation by removing either the isopropyl or the ethyl moiety of atrazine [182]. The same organisms carried out the dehalogenation, when incubated with both mono-N-dealkylated derivates, suggesting that such dechlorination

18 occurred a i ~ r the elimination of one or both alkyl groups (Figure 11). The mechanism of dechlorination of chloro-S-triazines, formerly attributed to chemical hydrolysis which occurs spontaneously in soil [175,183, 184], is now demonstrated to occur also in microorganisms, and this hydrolytic mechanism is responsible for the removal of the toxicity of these compounds. A new degradation product of atrazine in the soil was identified by Tafuri et al. as 2-isopropopylamino-4-methoxy-6methylamino-l,3,5-triazine [185]. However, the substitution o f - C 1 by OCH 8 seems to be a non biological process, and was in agreement with Pape and Zabik [186], who reported the formation of methoxy analogues of chloro-l,3,5-triazine herbicides when photolyzed in methanol under laboratory conditions in natural sunlight. Although anaerobic conditions may provide a much more favourable environment t h a n do aerobic conditions for hydrolytic dehalogenation which favour chloro-S-triazines degradation, the topic of anaerobic degradation of these compounds has not been widely studied till todate. Only Jessee et al. have reported the isolation of a facultative anaerobic bacterium which degrades atrazine under anaerobic conditions in a defined medium [187]. With regard to non-chlorinated triazines, such as ammelide, ammeline, melamine, cyanuric acid (CA) etc., a conclusive report for rapid and complete bacterial degradation was presented by Cook and Hiitter, who isolated

~i

I

~t

II

?H

v

H2N-- C . ~ ~-- NH-'-iC3H7 C! N Atrazine

ct -"iC3H 7 ~

~c

H2N--

N

oH

III H

iv

N,'~:~N ~.../ NH-- iC3H7 H2N--~JI~.l~S-.--

N,"'% N 2 HsC2"- I"IN--~LN~'~C--NH OH

Vl

N-,"C~N HaN--C ~ # C --'NH 2

Figure 11. Intermediates identified in the microbial degradation of atrazine: a, in Aspergillus fumigatus Fres. [176]; b, in Nocardia sp. [180]; c, in Pseudomonas sp. [182]. I, 2-chloro-4-amino-6-isopropylamino-S-triazine; II, 2-chloro-4-ethylamino-6-amino-S-triazine; III, 4-amino-2-chloro-Striazine; IV, 2-hydroxy-4-amino-6-isopropylamino-S-triazine; V, 2-hydroxy4-ethylamino-6-amino-S-triazine; VI, 2-hydroxy-4,6-diamino-S-triazine.

19 organisms, utilizing S-triazines as nitrogen source [188]. Melamine (triamino-S-triazine) is a compound, widely used as commercial chemical in the synthesis of industrial polymers. For this compound, Yutzi et al. proposed a degradative pathway with a Pseudomonas sp. strain A, involving three successive deaminations leading to the formation of CA, which is further metabolized [189]. The mechanism of each of the three deaminations and ring cleavage appears to be hydrolytic, as the reactions proceed in the absence of 02 . The four hydrolytic activities have been separated, exhibiting different characteristics: the reaction products from melamine were ammeline, ammelide, and CA. At each step, NI-I4* in equimolar amount was liberated. An analogous metabolic pathway was d e m o n s t r a t e d by Cook et al. for N-cyclopropylmelamine, a representative of the most highly aminated groups of mono-N-alkylated S-triazines [190]. This compound was shown to be degraded quantitatively to ammonium ion by two Pseudomonas spp. strains A and D. The first two reactions were quantitative stoichiometric deaminations catalyzed, exclusively, by strain A to give N-cyclopropylammelide. This intermediate was degraded exclusively by strain D in a quantitative stoichiometric hydrolytic dealkylamination process to CA, which is further converted into ammonium ion by both strains A and D. In this case, the alkylamino substituent is replaced b y - O H and not b y - H [180]. CA is the last S-triazine ring system in the degradation pathway of S-triazine herbicides. This compound is known to be utilized as the sole and growth-limiting source of nitrogen by bacteria [174]. Growth of a facultative anaerobic bacterium under anaerobic conditions on CA is reported by Jessee et al. [187]. Its degradative pathway was conclusively demonstrated by Beilstein and Hiitter in KZebsieUa pneumoniae [191], and by Cook et al. in a Pseudomonas sp. strain D [192] (Figure 12). OH

0 HN'~NH

~,~

__

H

H2N

NH 2 H

cyanuric acid

biuret

H20 CO2+NH H2O CO 2§

3 ~-- J

0

II ,,, H 2 N - - C - - N H

2

urea

Figure 12. Metabolic pathway of cyanuric acid in Pseudomonas sp. [192].

20 Crude extracts of strain D degraded CA, biuret, and urea quantitatively to NH4+ and CO 2 through three successive reactions, which occurred under either aerobic or anoxic conditions, and were presumed to be hydrolytic. From the results above reported, chlorinated and not S-triazines seem to be completely degraded to ammonia and carbon dioxide through different steps carried out, in some cases, by different organisms.

CARBAMATES,

PHENYLUREAS

AND

ANILIDES

Among pesticides and herbicides largely employed, carbamates and phenylureas play an important role, and their biotransformations are considered important for ecological and health reasons [193-195]. N-hydroxylated derivatives of carbamates have shown to be mutagenic [195]. Carbamates and phenylureas are derivatives of carbamic acid. While carbamates, also known as urethans, are esters, phenylureas are amides. Carbamic acid and N-substituted derivatives are unstable; they decompose spontaneously to carbon dioxide and ammonia or amine. Anilides are arylamides of carboxylic acids. For all these compounds, hydrolysis is the most important reaction involved in the biodegradation of ester or amide linkage (Figure 13) [196].

Ct A

l

Ct

II

N--C--C2Hs ~ ~ Propanil Clk):_~

B

~ H

Cl~

Linuron C[ ~~,, C

0

CH

I II / 3 N---C --N \

H 0 I II

Ct C[__~NH2

J

OCH3

CH3 i

Ct

N---C--O--CH I CH 3 Chlorpropham

Figure 13. Hydrolysis of herbicides: A, phenylamide; B, phenylurea; C, phenylcarbamate [196].

21 Carbamates This class of xenobiotics is s t r u c t u r a l l y and physiologically heterogeneous: Carbaryl and Carbofuran are acetylcholin esterase inhibitors, whereas Chlorpropham (CIPC) is a herbicide, interfering with cell division [197]. The structure of carbamates is very different, as the amine can be aromatic, like in CIPC, or aliphatic, like in Carbaryl and Carbofuran. Everywhere, these microorganisms are able to hydrolyze the carbamate linkage by producing CO 2, alcohol, and the corresponding amine, aromatic or aliphatic. Kaufman and Kearney have studied the degradation of several phenylcarbamates. Microorganisms, effective in degrading and utilizing these compounds as sole source of carbon, included Pseudomonas striata Chester, Flavobacterium sp., Achromobacter sp., Arthrobacter sp., and Agrobacterium sp. [198]. In the degradation of CIPC (isopropyl-N-3chlorophenyl-carbamate) , studied by utilizing an enzyme preparation from Pseudomonas sp., 3-chloroaniline was detected. This intermediate was derived from hydrolysis of CIPC; the same enzyme, however, was not able to hydrolyze 3-(p-chlorophenyl)-l, 1-dimethylurea (Monuron) [199]. In subsequent studies, Kearney reported purification and properties of this hydrolytic enzyme [200], and studied also the physico-chemical properties of twelve phenylcarbamate herbicides, in an attempt to determine their influence on microbial degradation. Studing the effects of the size of the alcoholic group on the rate of enzyme hydrolysis, Kearney found that by increasing the size of the alcoholic moiety, the rate of hydrolysis was retarded [201]. Whringht and Forey reported that Penicillium sp. hydrolyzed 4-chloro-2-butinyl-N-3-chlorophenyl carbamate) (Barban), yielding 3-chloroaniline [202]. Chloroanilines, derived from the degradation of chlorophenylcarbamates, are then degraded by several species of bacteria and fungi [203-205]. 2,3-Dihydro-2,2-dimethyl-7-benzofuranyl m e t h y l c a r b a m a t e (Carbofuran), extensively used to control corn rootworm, has shown to be slowly and incompletely degraded by soil microorganisms or fungi [206209]. Recently, it has been reported that an Achromobacter sp. WM 111, isolated by soil enrichment cultures, is capable of hydrolyzing Carbofuran at an exceptionally rapid rate. The organism utilizes the resulting methylamine as sole source of nitrogen, and produces 2,3-dihydro-2,2dimethyl-7-benzofuranol [210]. This microorganism catalyzes the degradation of other N-methylcarbamate insecticides (Carbaryl, Aldicarb, Baygon), but is ineffective in the degradation of acylanilide or urea pesticides. These results indicated that the hydrolytic enzyme of this strain is specific for phenol-carbamate ester linkages. In another report, the same authors by studying plasmid and chromosomal DNA of Achromobacter found that the strain harboured a plasmid which encoded for hydrolase activity [211]. Head et al. isolated, from soil, a bacterial strain, named MS2d which hydrolyzed Carbofuran, degrading phenolic

22 moiety. In addition to the observation that phenol degradation is mediated by a plasmid, initial evidence that a Carbofuran hydrolase gene is present on a second plasmid has also been given [212].

Phenylureas The decomposition of phenylurea herbicides by Bacillus sphaericus has been reported by Wallnvfer, [213], and Wallnvfer and Bader partially purified the enzyme responsible for hydrolytic cleavage [214]. Engelhardt et al. studied the degradation of (N-3,4-dichlorophenyl)-Nmethoxy-N-methyl urea (Linuron), and some other herbicides and fungicides with the same organism [215]. This microorganism degraded low levels of Linuron when it grew in the medium containing glucose, yeast extract, and asparagine, producing N,O-dimethylhydroxylamine, C02, and 3,4-dichloroaniline [216]. The amidase hydrolyzing Linuron was partially purified. This enzyme, named arylacylamidase, hydrolyzed also acylanilide herbicides and was also induced by different acylanilide herbicides, acylanilide fungicides and by phenylcarbamate herbicide (Propham); however, the maximum enzymatic activity was revealable with Linuron as inducer [217]. Diflubenzuron (1-(4-chlorophenyl)-3(2,6-difluoro-benzoyl)urea) was hydrolyzed in soil to 4-chlorophenylurea and 2,6-difluorobenzoate [218]. 4-Chlorophenylurea was presumably converted into 4-chloroaniline and bound to soil, while 2,6-difluorobenzoate mineralized to CO2 [219]. Phenylurea herbicides are also degraded under anaerobic conditions. N-(3,4-Dichlorophenyil)-N'-dimethylurea (Diuron) and Linuron have shown to be dechlorinated in anaerobic sediments with elimination of the chlorine atom in the para.position [220, 221].

Anilides Anilide herbicides constitute a group of compounds whose degradation occurs mainly through hydrolysis with formation of an acidic moiety, easily consumed by microorganisms. The herbicides N-(3,4-dichloro-phenyl) propionamide (Propanil), N-(3,4-dichlorophenyl) methylacryl amide (Dicryl), and N-(3,4-dichlorophenyl)-2-methylpentanamide (Karsil) were transformed into 3,4dichloroaniline [222-225]. The acylanilide hydrolases have been isolated from strains of Fusarium and Bacillus [226-228]. N-alkylacylanilides are, instead, compounds less easily hydrolyzable. The two most widely used are 2-chloro-2',6'- diethyl-N-(methoxymethyl)-acetanilide (Alachlor) and 2-chloro-2'-ethyl-6'-methyl-N-(1-methyl-2-methoxyethyl)-acetanilide (Metolachor). For these compounds, hydrolysis with arylacylamidases has not been reported as an important mechanism of degradation [229232], and they are very resistant to mineralisation. Novick et al. reported on the production of N-isopropylaniline from 2-chloro-Nisopropylacetanilide (Propachlor) by a microbial consortium [233].

23 Propachlor was also metabo|yzed by two microorganisms named DAK 3 and MAB 2 assigned respectively to the genera Moraxella and Xantobacter. DAK3 strain degraded Propachlor with the formation of catechol and 2-chloro-N-isopropyl-acetamide, which was released in the medium. MAB2 could grow on this metabolite (Figure 14). In this case, the metabolic pathway does not occur through a hydrolytic step [234].

O

C _N -cH2c' OAK3 II

OH Microbial biomass

H(CH3)2 Propachlor O II HN/C---CH2 CI \CH (CH3) 2

MAB2 .._

Microbial biomass

2-chloro-N i sop ropyl a r et amide

Figure 14. Proposed pathway of Propachlor degradation by strains DAK3 and MAB2 of MoraxeUa and Xanthobacter, respectively [234]. HALOGENATED

ANILINES

C h l o r i n a t e d a n i l i n e s are common m e t a b o l i t e s of t h e microbial degradation of various phenylurea, acylanilide and phenylcarbamates herbicides (Figure 13). Basicity and oxidability are the features of these compounds. They can react with oxygen and ozone present in air, with soil components [235-237], with lignin of plants [238], and with the microbial enzymatic systems. Frequently, they form condensation products. 3,4-Dichloroaniline may be transformed into 3,3',4,4'-tetrachloro-azobenzene [239-246] and different monochloro- and dichloroauilines are oxidised to dichloro- and tetrachloroazobenzenes [242,243]. An aniline oxidase and a peroxidase from the fungus Geotrichum candidum could dimerize different anilines except nitro anilines [244]. Condensation may transform anilines into nitroso benzenes [245]. Acetylation is an other reaction involved in the

24 m e t a b o l i s m of h a l o g e n a t e d anilines as detoxification m e c h a n i s m . 4-Chloroaniline was acetylated to 4-chloroacetanilide by bacteria [246, 247]; 4-amino-3,5-dichloroacetanilide was formed in soil by acetylation of 2,6-dichloro-l,4-diphenylendiamine, derived from the degradation of fungicide, 2,6-dichloro-4-nitroaniline [248,249]. Chlorinated anilines are not easily metabolized by microorganisms; however, cometabolic degradation of chloroanilines by several bacteria has been reported [250-256]. Both free and humus-bound chloroanilines were degraded at slow rates [235], but their mineralization was greatly enhanced by the presence of aniline [252]. A Rhodococcus strain converted 3-chloroaniline in presence of a growth substrated [254]. Zeyer and Kearney [257] and Zeyer et al. [258] isolated a Moraxella sp. strain, capable of utilizing chloroanilines and also 4-bromo- and 4-fluoroaniline, but not 4-iodoaniline as sole source of carbon, nitrogen, and energy. 4-Chloroaniline was metabolized by inducible oxygenase to 4-chlorocatechol, which was The aniline fttrther degraded via a modified ortho.cleavage pathway. oxygenase exhibited a broad substrate specificity. Some Pseudomonas acidovorans strains, with high degradative capacity toward, aniline, 3- and 4-chloroanilines to t h e c o r r e s p o n d i n g chlorocatechols, have been isolated [259]. This initial attack seems to be the limiting step in the rate of degradative process. Among the strains of Pseudomonas isolated, a Pseudomonas acidovorans CA2b also degraded slowly 2-chloroaniline. The three major reactions involved in the degradation of chloroanilines are summArised in Figure 15.

a/ NH2 / Ct

[~

N

C[

CI

NHCOCH3 CI

OH

~

CI Figure 15. Major aerobic biotransformation reactions of anilines: a, dimerization; b, acetylation; c, oxygenation.

halogenated

25 The anaerobic fate of chloroanilines has not been widely studied up to now. The reductive dehalogenation of chlorinated anilines has been investigated in methanogenic but non sulphate-reducing sediments by Kulm and Suflita and by Striujs and Rogers [260,261]. Methanogenic aquifer microbiota reductively dehalogenated di-, tri- and tetrachloroaniline by s e q u e n t i a l halogen r e m o v a l to t h e corresponding monoctdoroanilines which persisted for up to 8 months [262].

CONCLUSIONS Xenobiotic compounds have been used extensively in agriculture as herbicides and insecticides and in the m a n u f a c t u r i n g i n d u s t r y as surfactants, dyes, drugs, solvents and so on. Aliphatic and aromatic organic nitrogen compounds represent an important fraction of these chemicals. Even if many of the nitrogenous compounds are highly toxic and often recalcitrant to microbial attack, the microorganisms exposed to these synthetic chemicals have developed the ability to utilize some of them. L a b o r a t o r y s t u d i e s have e s t a b l i s h e d t h e c a p a c i t y of m a n y microorganisms to degrade or to t r a n s f o r m nitrogenous compounds under aerobic or anaerobic conditions. The results obtained from these investigations which involved pure cultures, or cell-flee extracts of microorganisms are important, because they may be predictive of their environmental fate. However, these studies do not reproduce the conditions found in nature where these organisms are exposed to a mixture of compounds and interact with other microbial communities. For every compound, which has proven to be biodegradable, the load of environmental pollutants is reduced. The assessment of biodegradability opens the way for the development of microbiological methods for the clean-up of soils and waters, contaminated with synthetic compounds. As bioremediation has its basis in the physiology and ecology of microorganisms, these methods have to be developed according to the capabilities of these microorganisms to ensure an optimal performance in those habitats. Moreover, the development of genetic manipulation techniques gives us the possibility to construct new strains with the desired "capabilities" for the degradation of xenobiotics. The employment of these strains could enhance the possibilities to decontaminate polluted environments.

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Biotransformations: Microbial Degradation of Health Risk Compounds Ved Pal Singh, editor 9 1995 Elsevier Science B.V. All rights reserved.

Synthesis and degradation of dimethyl the natural environment and in humans Tadashi

37 nitrosamine

in

Yoshinari

Wadsworth Center for Laboratories and Research, New York State Department of Health, and School of Public Health, State University of New York at Albany, Empire State Plaza, P.O. Box 509, Albany, NY 12201-0509, U.S.A.

INTRODUCTION Dimethyl nitrosamine (DMNA) is a simplest form of N-nitrosamines. DMNA and other nitrosamineS are carcinogenic [1,2] as well as teratogenic and m u t a g e n i c [3]. Since Magee and B a r n e s [4] discovered the carcinogenicity of DMNA with experimental animals, there have been numerous studies on DMNA and other nitrosamines in various aspects: mechanism of induction of carcinogenesis, factors promoting or suppressing production, mechanism of formation, and degradation in different environments, and distribution in food and other environments. Because of the enormous voh!me of reports of studies on nitrosamines, many of the recent reviews are focused on a narrow topic; endogenous N-nitrosation [5], bacterial mutagenesis by nitrosamines [6], occurrence of nitrosamines in foodstuffs, and dietary exposure to nitrosamines [7,8,9], environmental exposure to preformed nitroso compounds [10], and the levels of both N-nitroso compounds and their precursors in the human environment [11]. While bacterially mediated N-nitrosation has been reviewed as a part of endogenous N-nitrosation [5], there have been no comprehensive reviews on the production and degradation of DMNA in various environments, that are closely associated with microbial processes. In t h i s chapter, the processes of DMNA formation in various environments and the degradation of DMNA and other nitrosamines by various biogenic processes are reviewed. Endogenous formation and degradation of DMNA in humans are also included in this review, since they are involved with microbial processes in various degrees and are also considered to be the most important reactions that induce human cancer.

CHEMISTRY AND PRECITRSORS OF DMINA

Chemistry of DMNA DMNA and other nitrosamines can be formed chemically from nitrite and secondary amines (Figure 1). The mechanism of N-nitrosation is,

38

R

+

HO-NO

R,>N NO + H2 0

R

Figure 1. Formation of nitrosamines through N-nitrosation of secondary amines. however, much more complex [12, 13] than the reaction scheme shown in Figure 1. DMNA can be degraded by chemical or microbial processes or microsomal P-450 in liver microsomes. In all cases, the decomposition is either by denitrosation or demethylation (Figure 2). Denitrosation, which produces nitrite, can occur by both chemical and microbial reactions, while demethylation occurs by microbial and microsomal reactions. Sources of amines and nitrate Amines and nitrate are the precursors of DMNA. The following is a brief summary of the sources of these compounds. Amines are derived from food ingestion. Most amines, formed in the lower gastrointestinal tract, enter an enterogastric cycle prior to renal excretion. Mono-, di- and trimethylamines in h , m a n gastric fluid resemble those present in human saliva and blood [14]. Dimethylamine (DMA) and piperidine (both ~1 ~g/m]) are the most commonly occurring

H3C CH20H H3C OH3 N

H3C CH3 H3 C CH2 ~N/'+ "N/

e"

~

0

+

H

0

/

A A)

OH3

,,B ~ "hi ~0 HCHO "

~\(B) H 3C\ C H 2 - ~ N

~-~

- ,NO~N02 1_ 02

.,~. N2

3

HCHO + H3C-NH2 ~,,

~NO~

2

Figure 2. Possible mechanisms of the DMNA metabolism (modified from the schemes proposed by Wade et al. [80]) by demethylation (A) and denitrosation (B).

39 nitrosatable secondary amines in h u m a n saliva, gastric juice, blood, urine, and faeces [15]. In a typical dietary intake of 7.5 rag/day of total secondary amines, dimethyl~mine constitutes as much as 59% [16]. Nitrate, being a precursor of D M N A and other nitrosamines, is a significant risk factor in h u m a n cancer [17,18,19]. Amongst all, nitrate of dietary and dmnking-water origin [20] is a major source of endogenous nitrosation, and is actively absorbed in the small intestine, circulated in the bloodstream, and secreted into saliva [21]. Salivary nitrate is reduced to nitrite by bacteria, containing nitrate reductase enzymes, while oral nitrate reduction represents 80% of an individual's nitrite exposure [22]. These bacteria colonize h u m a n saliva at levels averaging 107 CFU/ml [23]. The nitrite level of h u m a n saliva is correlated with the amount of dietary nitrate [24,25]. It should be noted that the life of nitrite in the bloodstream is very short, primarily owing to scavenging by oxyhemoglobin [26,27]. Nitrite in saliva transported to the stomach will become an effective nitrosating agent in the stomach, where the p H is low and secondary and tertiary amines are present. In fact, the existence of a quantitative relationship between oral nitrate-reducing activity and the endogenous formation of N-nitrosoamino acids in humans has been demonstrated [28]. The level of oral nitrate reduction appears to be the major factor affecting the gastric nitrosation [28]. Another important source of endogenous nitrate in humans is biosynthesis from L-arginine [29-32]. The range of daily endogenous biosynthesis of nitrate is 1-1.2 retool.person-I [33,34]. Determining the incorporation of 15NHs, Wagner et al. [35] found that nitrate biosynthesis was enhanced by endotoxin treatment. Stuehr and Marletta [36] found synthesis of nitrite and nitrate in routine macrophage cell lines. While nitrite, nitrate, nitric oxide, and N-nitrosating agents are known to be synthesized by some m a m m a l i a n cells [37], it was found recently that nitric oxide synthase activity can be induced by toxic shock syndrome toxin 1 in a macrophage-monocyte cell line [38]. FORMATION

OF

DMNA

The formation of DMNA and other nitros~mines by microbial reactions may also be represented by the reaction scheme shown in Figure 1. Much of the efforts, to find the distribution, formation, and degradation of DMNA in different natural environments and by various microbes, were made in the 1970s. The main reason, that there are not many reports on any of those aspects in recent years, may be because the impact is not as great as the other routes of carcinogenesis of humans, as is discussed in later section.

40 F o r m a t i o n in the natural e n v i r o n m e n t DMNA can be formed by biochemical processes in various natural e n v i r o n m e n t s . Factors, affecting the formation and s t a b i l i t y of nitrosamines in different environments, were initially investigated by Alexander and his associates. For example, Ayanaba et al. [39] studied the possible microbial contribution to nitrosamine formation in sewage and soil. Subsequently, transformation of methylamines and formation of DMNA in samples of treated sewage and lake water [40], stability of nitrosamines in samples of lake water, soil, and sewage [41], and factors affecting DMNA formation in soils and water [42] were examined. Kobayashi et al. [43] found that DMNA can be formed in a polluted environment. They also confirmed that DMNA is formed in artificial wastewater in the range of pH 4-9 in the presence of nitrite. However, there is little evidence to support the view that the production of DMNA and other nitrosAmines by microbial actions is an important process in natural environments [44]. From their studies with cultures of several microorganisms, Mills and Alexander [45] found that Pseudomonas stutzeri was able to catalyze nitrosation, only when in the growing phase. Although the species was not identified, Ishiwata et al. [46,47] found that bacteria in human saliva formed DMNA from nitrate and dimethylamine. Microbial formation There are a large number of reports on DMNA formation from sodium nitrite and dimethylamine by enteric bacteria [48-53], and by resting cells of Escherichia coli B [54]. Using a sensitive fluorimetric method to determine nitrosamines, Ralt et al. [55] showed that nitrosation by E. coli can be induced under anaerobic conditions by nitrite and nitrate. Ji and Hollocher [56] found that the nitrosation reaction by E. coli was carried out first by the production of NO from nitrite, followed by O2-dependent chemical nitrosation. The nitrosation is a chemical reaction, that proceeds with N208 and N204 derived from NO. E n d o g e n o u s f o r m a t i o n of DMNA Endogenous nitrosation is a process that is involved with the formation of DMNA and other nitrosamines in human body, with or without the involvement of microbial activity. The reaction can take place in different parts of the ht~man body. Since a strong correlation appears between the exposure of humans to endogenously formed N-nitroso compounds and the induction of cancer [57], extensive studies have been in progress in recent years. Endogenous N-nitrosation may result from cell-mediated reactions, involving alveolar and peritoneal macrophages, bacteria, and yeasts [5,58,59]. In earlier investigation, dimethy|amine was used to determine

41 the rate of nitrosation of amine [52,54]. In contrast, a series of studies were made with morpholine by Calmes et al. [60-62] because the rate of nitrosation of morpholine was about 30 times faster t h a n t h a t of dimethylamine. While the maximum rate of DMNA formation by resting cells of Escherichia coli was observed at pH 8.0 [54], Calmels et al. [60] found several species of microorganisms that are capable of nitrosation catalysis at varied rates at pH 7.25. They also studied kinetics of nitrosamlne formation from secondary amines by E. coli strains. In this paper they exAmlned substrate specificity of three E. coli strains to catalyze nitrosation of various amines. On the basis of their kinetic studies for nitrosation from secondary amines by E. coli A10, Calmels et al. [61] concluded that bacterial nitrosation is an enzyme-mediated reaction, closely associated with molybdenoenzymes, such as the nitrate reductase/formate hydrogenase system. Subsequent reports [55,62] confirmed the involvement of nitrate reductase in nitrosation. The comparison of relative activity between chemical and bacterial nitrosamine formation appropriate to gastric juice was made [58]. Nitrosation of amines can be stimulated by the presence of macrophages [63]. D M N A can be formed at neutral pH (achlorhydric gastric juice and infected organs, such as bladder) with assistance from microbial activity (nitrosating enzymes). Bacterially catalyzed N-nitrosation reactions proceed much more rapidly at neutral pH than the chemical reaction [58]. Among the species and different strains of bacteria, Leach et al. [58] found that the most rapid catalysis was associated with those bacteria, capable of reducing nitrate and nitrite by the process of denitrification. Their study was based on the use of Pseudomonas aeruginosa BM1030, an isolate from achlorhydric gastric juice. This is in contrast to the conclusion by Licht and Deen [64], who have developed a mathematical model to estimate the rate of formation of nitrosamine in human stomach. The calculated amount of gastric formation of DMNA was ~0.02 nmol/ day. As the amount is a factor of ~10 ~ to 108 lower than published estimates of dietary exposure to preformed DMNA, gastric formation of DMNA does not pose a serious additional health risk so long as the stomach is acidic. However, in achlorhydric stomach it may be a serious t h r e a t of carcinogenesis due to increased production of nitrosamines. Increased levels of nitrosamines in blood of patients, with chronic renal failure [65], in urine of patients with infection of the bladder due to urinary diversions [10], and in the urinary tract of paraplegic patients [9] indicate t h a t DMNA was formed endogenously by a bacterially mediated in vivo formation of N-nitroso compounds. Calmels et al. [66] isolated an enzyme catalyzing nitrosamine formation in Pseudomonas aeruginosa and Neisseria mucosae at n e u t r a l pH. Their results suggest t h a t carcinogenic N-nitroso compounds may be formed endogenously with nitrosating enzymes t h a t are provided by some bacteria.

42 As vitamin E acts as nitrite scavenger, it serves as inhibitor of the formation of N-nitrosamine [67].

DEGRADATION

OF

DMNA

There are two types of microbial degradation of D M N A (Pathways A and B in Figure 2). One is denitrosation, which produces nitrite and amine. This is a reverse of the process of chemical formation of D M N A (Figure 1), and is presumed to be a main process for removal of D M N A from the environment. The other is demethylation, which produces aldehyde and methyl amine. This reaction is considered as a central paradigm for initiation of carcinogenesis. Natural environment Tate and Alexander [68] studied resistance of nitrosamines to microbial attack in the environment. Degradation of nitrosamines in lake water and sewage [69] and in the marine environment [70] has been reported. Kaplan and Kaplan [71] reported biodegradation of DMNA in aqueous and soil systems. In both systems, the rate of degradation was slow, but linearly correlated with the concentration of DMNA (Figure 3). On the

100 ng ~.

1 mg

A

i

cO "O

"7

.,.,,,

E

'

10

1

ng

pg 10 ~g

r 0 loong

..~

100 fg

N L_

(%) c

1 ng

10 ng

.......

1 gg

,

,

100 I.Ig

.....

DMNA (g soil -1)

10 mg

fQ

i

pg

i

100 pg

..L

.

A

.

!

10 ng

DMNA

l

ling

l_

__

100 pg

(ml-')

Figure 3. Initial rates of mineralization over a range of concentrations of D M N A in (A) soil batch culture and (B) aqueous batch culture. Symbols:O, lake water with salts;O, lake water;A, lakewater with salts and glucose [71].

43 basis of their finding, that the substrate concentration reduction curves generated with DMNA were not sigmoidal, the possibility of the DMNA metabolism by the biomass being growth-related was ruled out. Although the characteristics of the microbes are not known, they identified two metabolic intermediates, formaldehyde and methylamine, from the DMNA degradation. Pure culture studies Five out of 44 species of bacteria, molds, and yeasts were found to form nitrite from DMNA [72]. Using Rhizopus oryzae, Streptococcus cremoris, and Saccharomyces rouxii, Harada and Yamada [73] examined the degradation of five different N-nitrosamines, including DMNA. While N-nitrosodipropylamine (DPNA) was most easily degradable, DMNA was least degraded among five different nitrosamines. Their results with whole-cell cultures suggested that the enzymes responsible for the degradation of N-nitrosodipropylamine were inducible. Using whole cells in growth phase and cell-free extract of Rhizopus oryzae, Harada [74] studied further the microbial degradation of DMNA and DPNA. The degradation activities in the cell-free extract were highest at 30~ and pH 8.0, under anaerobic conditions. The degradation of DMNA and DPNA was concentration-dependent with cell-free extract, while that with whole cells in the growth phase showed a maximum at -0.1 raM. A series of these studies did not investigate the metabolic pathways of DMNA_ Kobayashi et al. [43] found that photosynthetic bacterium, Rhodopseudomonas capsulata, is capable of metabolizing DMNA. Various enteric bacteria metabolized DMNA through denitrosation, by which dimethylamine and nitrite were formed, and dimethylamine was not further metabolized [75]. The other type of the degradation of DMNA is by demethylation. It was found that a methanotroph, Methylosinus trichosporium OB3b (MT OB3b), degraded DMNA in the presence of methane, presumably by the catalytic action of NAD(P)H-dependent methane monooxygenase (MMO) [76]. Tracer studies with 14C-labelled DMNA revealed that MT OB3b was capable of both assimilating DMNA-carbon into the cell and respiring it as CO 2 (Figure 4). The rates of CO 2 production (Vco2) from and cellular u p t a k e (Vp) of DMNA were linearly correlated with the DMNA concentrations of 0.03-10 mM, which corresponded to approximately 3 per cent of the added DMNA metabolized in 24 h. These rates were two to three orders of magnitude less than that of the uptake of methane (VcH4). VCH4 was suppressed only when the concentrations of DMNA exceeded 0.3 mM (Figure 4). In the presence of 0.1 mM DMNA, Vp and Vco, were essentially the same with or without the presence of methane in the first 8 h of incubation, but declined sharply thereafter only when methane was absent. These observations suggested that the metabolism

44

100

- ,,t,,,-,!

IE

~" z

J

10

Y

J

17

1

a

-~

0.1

E e--

9

:

:

~

9:

:

:

;

:

,

;

'

,

e I

--

i

IE

C" r

==,=.,

0

IE

>o

4 2 1

I

"---.o

'3-

0 . I , "

o .01

;

;

;

'

I

0.1

:

DMNA

;

:

:

[

1.0

:

'"'

'

;

I

10

(mM)

Figure 4. Metabolism of DMNA by Methylosinus trichosporium OB3b (MT OB3b) in the presence of different concentrations of DMNA. Average rates at 0-24 h of incubation are given [76]. The rates of cellular uptake, CO~ production, and methane metabolism are represented by Vp (Q), Vco 2 (M) and VcH4 (O), respectively.

of D M N A was carried out by methane monooxygenase (MMO), and that N A D H , a cofactor for M M O , m a y be provided from the oxidation of the stored energy in the cells, when methane is not available. The sequence of the reaction of D M N A by M T OB3b has not been clearly understood. However, in view of the finding that the D M N A carbon was incorporated to cell material and respired as C O 2, Yoshinari and Sharer [76] postulated that M T OB3b produced formaldehyde through

45

CELL CONSTITUENTS

MMO CH4 ~ NADH2

- CH3OH,/.-~..~ HCHO..,,~..--,..,~ H C O O H ~ NAD+

X

XH2

NAD+ NADH2

- CO2 +H2 0

NAD+ NADH2

Figure 5. Schematic mathane oxidation pathway by methanotrophs. Initial step for the metabolism of methane is catalyzed by methane monooxygenase (MMO).

hydroxylation of D M N A by M M O , as is the case with the oxidation of methane (Figure 5). They suggested that the initial step for the formation of formaldehyde could be quite similar to the reaction sequence carried out by a liver microsomal cytochrome P-450-dependent mixed-function oxidase (MFO). Both M M O and M F O require cofactor N A D ( P ) H and oxygen for their activity. With M F O , DMNA is activated to a-hydroxylnitrosamine. Subsequently, it is decomposed to form spontaneously formaldehyde and methyldiazonium (Pathway A in Figure 2). Most of the unstable reactive intermediate, CHzN2, which can interact with DNA and other nucleophiles for carcinogenicity and mutagenicity, is suspected to react with water and form ROH and N 2. To test further whether MT OB3b metabolizes DMNA by this sequence, it would be of interest to determine the production of ~sN2 from the lSN-labelled DMNA, by using the method of Kroeger-Koepke et al. [77].

Endogenous

degradation

of DMNA

A large body of information regarding metabolism of DMNA by m a m m a l i a n cells is documented. Mixed-function cytochrome P-450 isozymes (MFO) in mammalian liver microsomes are involved in the metabolism of DMNA [1,6,78]. The same cytochrome P-450 isozymes are reported to be involved in both nitrite formation (denitrosation) and a-hydroxylation (demethylation) of nitrosamines [79-81].

Denitrosation Denitrosation results in the detoxification of DMNA. Keefer et al. [82] found concurrent generation of methylamine and nitrite during

46 denitrosation of DMNA by rat liver microsomes. Amelizad et al. [83] studied the effect of antibodies against cytochrome P-450 on demethylation and d e n i t r o s a t i o n of DMNA and N-nitrosomethylaniline. Hydroxy derivatives, aldehydes, and nitrite were formed from N-nitrosomethylN.amylamine by rat liver microsomes and by purified cytochrome P-450 IIB1 [84]. Recently, an increased oxidation of DMNA in pericentral microsomes, after pyrazole induction of cytochrome P-450 2El, has been reported [85]. Heur et al. [86] studied the Fenton degradation as a nonenzymic model for microsomal denitrosation of DMNA.

Demethylation This reaction is the central paradigm for initiation of carcinogenesis. DMNA is metabolically activated to generate the ultimate carcinogenic form. Initial step in this biotransformation is believed to be enzymatic hydroxylation by MFO, followed by spontaneous cleavage of the carbonnitrogen bond, which releases aldehyde and methyldiazonium (Figure 2). Most of the unstable intermediate, CH2N2, that is suspected to interact with DNA and other nucleophiles for carcinogenicity and mutagenicity, seems to react with water and form methanol and N r Methyl diazohydroxide appears to be the main cause of carcinogenesis and mutagenesis [1,6,78]. Jensen et al. [87] provided evidence that the in vitro methylation of DNA was carried out by microsomally-activated DMNA and that the methylation was correlated with formaldehyde production. Sagelsdorff et al. [88] showed DNA methylation in rat liver by daminozide, 1, 1dimethylhydrazine, and DMNA. Studies with other nitroso compounds also confirmed the formation of hydroxy derivatives, aldehydes, and nitrite by rat liver microsomes and by purified cytochrome P-450 IIB1 [84]. The sequence of specific methylation of DNA by N-nitroso compounds shares a common intermediate, methyl diazonium ion [89]. CONCLUSIONS Dimethyl nitrosamine (DMNA) is a potent carcinogenic compound. Unlike the xenobiotic compounds, that are known to be carcinogenic, it is not the product of industrial processes. Instead, DMNA is the product of chemical and microbial processes in the natural environment and in the human body system. A major source of DMNA for humans is endogenous production rather than ingestion from the food. DMNA can be decomposed chemically and biochemically. The product derived from the enzymatic degradation of DMNA is responsible for methylating the nucleotides, that trigger the process of carcinogenesis.

47 Endogenously formed DMNA in humans through chemically and/or bacterially mediated reactions, but not of natural environmental origin, appears to be mainly responsible for causing human cancer. There are two pathways of degradation of DMNA by microbial enzymes and by microsomal P-450. One is denitrosation and the other demethylation. Denitrosation is a main microbial process for removal of DMNA from the environment, although its degradation through demethylation also takes place by a methanotroph. The reaction catalyzed by microsomal P-450 can couple with the methylation of DNA, a central paradigm for initiation of carcinogenesis.

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1 2 3

4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

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Yordy JR, Alexauder M. Appl Environ Microbiol 1980; 39: 559-565. Petit L, Aubert J, Dubreuil A, Levrat B. Rev Int Oceanogr Med 1984; 73-74: 41-50. Kaplau DL, I~plau AM. Environ Microbiol 1985; 50: 1077-1086. Harada K, Yamada I~ J Shimonoseki Univ Fish 1978; 26: 293-298. Harada K, Yamada I~ Nippon Suisan Gakkaishi 1978; 45: 925928. Harada I~ Bull Jap Soc Sci Fish 1980; 46: 723-726. Rowland IR, Grasso P. Applied Microbiol 1975; 29: 7-12. Yoshinari T, Shafer D. Can J Microbiol 1990; 36: 834-838. Kroeger-Koepke MB, Koepke SR, McClusky GA, Magee PN, Michejda CJ. Proc Natl Acad Sci USA 1981; 78: 6489-6493. Magee PN, Barnes JM. Adv Cancer Res 1967; 10: 163-246. Tu YY, Yang CS. Arch Biochem Biophys 1985; 242: 32-40. Wade D, Yang CS, Metral CJ, Roman JM, Hrabie JA, Riggs CW, Anjo T, Keefer LK, Mico BA. Cancer Res 1987; 47: 3373-3377. Sohn OS, Ishizaki H, Yang CS, Fiala ES. Carcinogenesis 1991; 12: 127-131. Keefer LK, Anjo T, Wade D, Wang T, Yang CS. Cancer Res 1987; 47: 447-452. Amelizad S, Appel KE, Oesch F, Hildebrandt AG. J Cancer Res Clin Oncol 1988; 114: 380-384. Ji C, Mirvish SS, Nickols J, Ishizaki H, Lee MJ, Yang CS. Cancer Res 1989; 49: 5299-5304. Dicker E, Cederbaum AI. Clin Exp Res 1991; 116: 1072-1076. Heur YH, Streeter AJ, Nims RW, Keefer LI~ Chem Res Toxicol 1989; 2: 247-253. Jensen DE, Lotlikar PD, Magee PM. Carcinogenesis 1981; 2: 349354. Sagelsdorff P, Lutz WK, Schlatter C. Fund Appl Toxicol 1988; 11: 723-730. Milligan JR, Hirani-Hojatti S, Catz-Biro L, Archer MC. Chem-Biol Interactions 1989; 72: 175-189.

Biotransformations: Microbial Degradation of Health Risk Compounds Ved Pal Singh, editor 9 1995 Elsevier Science B.V. All rights reserved.

Aflatoxin

biotransformations:

51

biodetoxification

aspects

Ved Pal Singh Department

of Botany,

University

of Delhi,

Delhi-110007,

India

INTRODUCTION Since antiquity, man has been witnessing the hazardous effects of mycotoxins on h u m a n and animal health, as almost all agricultural produce, including most foods and feeds used for h u m a n and animal consumption, get contaminated by the toxigenic fungi. Mycotoxins are highly poisonous secondary metabolites produced mainly by filamentous fungi [1], which contaminate the foods and feeds at some stage of their production, processing, transportation or storage. Over 200 mycotoxins have already been identified [2], and still many others are yet to be characterized. Of all mycotoxins, aflatoxins constitute the most widely studied, and their toxic, mutagenic as well as carcinogenic effects on both man and animals are weU-documented [1,3-12]. Aflatoxins are also known to influence metabolism of higher plants, ferns, algae, fungi, and bacteria [13]. Aflatoxins were first discovered by Sargeant and coworkers in England, in the year 1961, while investigating the causal factors for widespread episode of unexplained mortality encountered in poultry flocks, swine, and cattle due to a disease which later came to be known as the "Turkey. X' disease. These toxins, found in mould-infected peanut meals, were responsible for the Turkey-X disease in livestock. The mould was identified at the Royal Botanic Gardens, Kew, as Aspergillus flavus, and the corresponding toxin was named after this fungus as A. flavus toxin or "Aflatoxin'. These potentially h a z a r d o u s secondary metabolites are produced primarily by some strains of A. flavus, A. versicolor, A. nidulans and most, if not all, strains of A. parasiticus as well as a related species A. nomius [14]. Aflatoxigenic fungi are more commonly found in tropical and subtropical areas where both temperature and humidity are favourable for growth. Aflatoxins are highly oxygenated, heterocyclic compounds, having dihydrofurano or tetrahydrofurano moieties fused to a substituted coumarine moiety and are synthesized through polyketide pathway, linking both primary metabolism, i.e., biosynthesis of fatty acids and secondary metabolism (polyketide biosynthesis), involving an intermediate (acetyl coenzyme A) between these pathways for aflatcoxin biosynthesis. Aflatoxins have been characterized on the basis of their fluorescence properties. Those which fluoresce blue under UV light are classified as B aflatoxins and those with green fluorescence are classified as G aflatoxins. And on the basis of their relative mobilities (Rf values) on TLC plates, these toxins are further classified as aflatoxins B~ and B 2 (AFB 1 and AFB 2) and aflatoxins G 1 and G 2 (AFG 1 and AFG2).

52 That aflatoxins are acutely toxic to m a n was demonstrated by an outbreak in India in 1974 of epidemic jaundice, involving severe liver disease, which resulted in the deaths of more than 100 people and serious illness in 400 others [15]. Aflatoxms B~ and G 1 have been shown to be mutagenic in bacteria, vegetative cells as well as in Drosophila melanogaster and Salmonella typhimurium. Also, metabolites have been studied for their mutagenic effects in S. typhimurium and show large variations in their mutagenic potency [16]. Carcinogenicity of aflatoxins has been demonstrated in various animal species [3,12,17,18]. A F B 1 induces malignant tumours in rats, mice, monkeys, marmosets, ducks, guppies, salmon, trout, and tree shrews. The liver is the target organ for these compounds, but some pulmonary turnouts have been observed in treated mice, as well as kidney and intestinal turnouts in rats. AFB~ produces carcinogenic effects in m a n y species, following exposures to low doses (as low as 1 ~tg/kg in the diet). Aflatoxins must be considered to be a probable causative agent for primary liver cancer in man, which is endemic to Thailand, Kenya, Switzerland, and Mozambique. AFG~ and A F B 2 are also known to induce liver turnouts in some animals [18]. Considering all these aspects of aflatoxin-induced health hazards in m a n and animals, and because of the widespread occurrence of these toxins in otherwise nutritious natural products, m a n y studies have been carried out to find effective and suitable as well as convenient methods of detoxification of aflatoxins. P H Y S I C A L AND CHF_J~CAL DETOXIFICATION

METHODS

OF

AFLATOXIN

Although physical and chemical methods of detoxification are not the subject m a t t e r of the present review, some recent works have been included to have a latest insight into the mechanism of aflatoxin detoxification. Some of the conventional methods have already been mentioned elsewhere [19]. Effects of physical factors, such as light, temperature, pH, etc., on aflatoxin detoxification have been of interest to biotechnologists and food scientists. The reduced levels of ailatoxins in contom~nated grains of rice due to green light and in those of ragi as well as in liquid broth due to blue light have been recorded by Shrivastava et al. [20]. In artificially infected corn meal and peanuts with Aspergillus parasiticus, chlorine gas has been found to show 75% degradation of AFB 1 [21]. The mutagenicity of chlorine-treated copra meals and peanuts spiked with AFB1 was greatly reduced compared with untreated controls, as determined in Salmonella typhimurium strain TA98 in the presence of rat liver

53 5-9 mix; the decrease in the mutagenic potential has been correlated with reduction in AFB~ levels. However, no mutagenic compound was generated by such treatment. Similar results have been obtained with ammonia treatment of aflatoxin-contaminated cotton seeds by Jorgensen et al. [22]. Abdel-Rahim et al. [23] have shown that the rate of detoxification of aflatoxins in cotton seeds increased with increasing concentrations of ammonil!m hyroxide (NH4OH) up to 0.4%, then decreased at further higher concentrations. The detoxiiication rate for AFB~ was higher than that of AFB 2 and AFG 2 at very low concentration (0.1%) of NH4OH. Aqua-ammonia method has been quite successful in detoxifying the aflatoxins in contaminated poultry feed to a non-detectable level on 3rd day after ammoniation, using 1.5% aqueous ammonia solution sprayed over contaminated feed [24]. Interestingly enough, a combination of physical and chemical (physicochemical) m e t h o d h a s been quite effective in AFB~ d e g r a d a t i o n . Degradation of aflatoxin B1 in dried figs by sodium bisulphite with or without heat, UV energy or H202 has been studied by Altug et al. [25]. There was 28.2% degradation by 1% sodium bisulphite treatment alone, but when H202 (0.2%) was added, 10 rain before bisulphite treatment, 65.5% degradation of AFB~ was achieved in 72 h. In both cases, maximum degradation occurred during 2nd day of treatment. Heating of the bisulphite-treated samples at 45~ to 65~ caused 68.2% degradation of AFB~. However, UV radiation degraded 45.7% of AFB~ in fig samples, when exposed for 30 mln, but the rate of degradation was not affected by the addition of bisulphite or H202.

BIODETOXIFICATION

OF

AFLATOXINS

Biodetoxification aspects can be further divided into three categories, depending on the type of system involved, which are as follows: (a) C o m m o d i t y - d e p e n d e n t detoxification. (b) E n z y m a t i c detoxification. (c) Microbial detoxification.

(a)

Commodity-Dependent

Detoxification

Plants can carry out biotransformations of aflatoxins, as made clear by Howes et al. [26], who studied the metabolism of aflatoxin B 1 in Petroselinum crispum (Parsley). On administration of AFB1 to whole plants, aflatoxicol was formed. The cell-free preparations, on the other hand, formed two new aflatoxin B~ or aflatoxicol A. Even plant products, such as neem-leaf extracts, have been shown to control aflatoxin production in Aspergillus flavus-infected cotton bolls [27]. Sometimes, the genetic constitution of the aflatoxin contaminated plant commodity can take

54 care of the levels of aflatoxins in itself [28]. For example, the tannin extracts from some genotypes of peanut cultivars, such as PI 337409 and TX 798736, have been found to reduce the level of aflatoxin production [29]. Also, certain corn genotypes which get contaminated with the toxigenic fungus, such as Aspergillus flavus, can regulate the toxin production in the grain [28], and this might be the reason why some genotypes of maize germplasm develop resistance to infection by A. flavus, and/or subsequent contamination by AFB 1 [30]. Not only the genotypes, but also the biomolecules of the plants, such as glyceollin of soybean seeds, have been implicated in lowering down the aflatoxin levels in the A. flavus-infected viable seeds [31]. On the other hand, Rasic et al. [32] reported t h a t the level of AFB~, added to a food commodity, i.e., milk, before fermentation at concentrations of 600, 1000 and 1400 ~g/kg, was reduced in yoghurts (pH 4.0) by 97, 91 and 90%, respectively. According to them, a decrease of AFB 1 (conc. 1000 /~g/ kg) in milk, acidified with citric, lactic, and acetic acids (pH 4.0), was 90, 84 and 73%, respectively.

(b) Enzymatic

Detoxification

Almost all human and animal species get exposed to aflatoxin doses beyond permissible limits, when they consume the aflatoxin-contaminated foods and feeds. However, there are special mechanisms by which aflatoxins are detoxified within the living systems, so as to eliminate the possibility of health hazards caused due to consumption of aflatoxinc o n t a m i n a t e d foods and feeds. It has been d e m o n s t r a t e d t h a t biotransformations of aflatoxins are mediated by microsomal enzymes, exhibiting the mixed function hydroxylases of endoplasmic reticulum in higher organisms [33]. The liver microsomal preparations from many animal species have been shown to transform the most hepatotoxic and hepatocarcinogenic aflatoxin - AFB 1 to a 4-hydroxy derivative (i.e., aflatoxin M 1, AFM 1) and an analogous product. Aflatoxin GM~ may be formed from AFG~. Both A F B 1 and AFG~ have been shown to be metabolized to their respective 2-hydroxy derivatives or hemiacetals by NADPH 2dependent enzyme [33]. Patterson [33] also reported the enzymatic detoxification of aflatoxins in the livers of rabbit, duckling, guinea-pig, chick, and mouse, following a minor route to the formation AFM~. Human liver microsomes were able to detoxify AFB 1 and convert it to AFQ~. However, once produced, AFQ~ was not appreciably oxidised in human liver microsomes and was not very genotoxic. The 3~-hydroxylation of AFB~ to AFQ1 is considered to be a potentially significant detoxification pathway [34]. Studies have also shown that mice become resistant to carcinogenic effects of aflatoxin B~ and that, this is due to expression of an isoenzyme of glutathione S:transferase (GST) with high activity towards AFBF8-9-epoxide [3]. Daniels et al. [35] have also demonstrated the biotransformations of potentially hepatotoxic and hepatocarcinogenic

55 AFB~ to less toxic metabolites, AFMz and AFQ1 in rabbit lung and liver microsomes. Various products of aflatoxin B 1 metabolism are given in Figure 1.

HO H H

H

HO~O H~~a,~

+

HO

A~ATOXICOL

~I REDUCTO IN HYDjOXYLAT(~ I N('~~

DEMETY i LATION EPOXYH

3-HYDROXY-

(71

OH

OH

Figure 1. Metabolic biotransformation products of aflatoxin B r Microbial systems show similarities to the animal systems, and Hamid and Smith [36] have clearly demonstrated the involvement of a microsomal enzyme system in aflatoxin degradation in A. flavus. The degradation of aflatoxins by cell-flee extracts of A. flavus was enhanced by NADPH [36], which is consistent with the activity of enzymes in which this cofactor is necessary for enhanced aflatoxin detoxification in eukaryotic systems [33]. NADPH-dependent 17-hydroxy-steroid dehydrogenase has been reported to transform aflatoxin B~ to aflatoxin R 0. Doyle and Marth [37] have observed maximum aflatoxin degradation by the intramycelial factors of the fungus at physiologically optimal pH and temperature, conducive for the aflatoxin-degrading enzymes.

56 Also, the enzyme which uses hydrogen peroxide (H202) aS the substrate appears, to play a key role in aflatoxin degradation or detoxification, as H202 enhances the aflatoxin degradative activity, when it was added to the mycelial proteins of A. parasiticus [6]. Thus, peroxidase could be the probable enzyme, which might possibly help in detoxification of aflatoxins. The studies of the author in collaboration with Professor John E. Smith at the University of Strathclyde, Glasgow (U.I~), have indicated the involvement of microsomal peroxidase in aflatoxin detoxiiication, without precluding the possibility of involvement of cytochrome P-450 monooxygenase in aflatoxin detoxification [36]. The role of hepatic microsomal P-450 monooxygenase in AFB 1 detoxification in animal systems has already been established. The studies on cytochromemediated metabolism of endogenous substrates, such as steroids and fatty acids as well as biotransformation of xenobiotics, have been welldocumented [38,39].

(c) Microbial Detoxification This aspect includes the involvement of microorganisms to detoxify aflatoxins. On the basis of the types of microorganisms involved and the methods through which they detoxify aflatoxins, microbial detoxification approaches can be grouped into the following three categories" (i) (i i)

Aflatoxin detoxification by the toxigenic fungus itself. Aflatoxin detoxiilcation by the atoxigenic strains of the same fungal species. (iii) Aflatoxin detoxification by other atoxigenic microorganisms.

(i) Aflatoxin detoxification by the toxigenic fungus itself Aflatoxin-producing fungi, such as A. flavus and A. parasiticus, are ubiquitous in nature and contaminate most foods and feeds, rendering them unsuitable for consumption (by producing aflatoxins) and posing potential threat to human and animal health. It would, indeed, be worthwhile to get these toxins detoxified by the producer organism itself by altering growth conditions, including change in pH and temperature, so as to enable it to detoxify its own toxin under such manipulated conditions. Studies have been carried out to demonstrate the ability of ~ flavus and A. parasiticus (both toxigenic) to degrade aflatoxins produced by themselves [6,36,37,40-45]. How this aflatoxin biotransformation is influenced by cultural conditions, such as growth substrates, age of culture, aeration, pH, and temperature, will be considered here briefly.

57

Effect of growth substrates, age of culture, and aeration aflatoxin detoxification

on

The growth substrates (either liquid or solid), used to produce mycelia, can affect the ability of the producer strains of fungi to degrade aflatoxins [42,44]. Marth and Doyle [44] have demonstrated that 9-day-old mycelia of A. parasiticus NRRL 2999, grown in potato dextrose, Czapek-Dox, and YES broths, exhibited little or no degradation of aflatoxin B~, while mycelia grown in glucose salt, Y-M and Moyer's broths were able to show increased levels of aflatoxin detoxification [44]. Shih and Marth [46] reported that aflatoxins B~, B2, G1, and G2 were degraded by 8- and 16-day-old, but not by 4-day-old mycelium of A. parasiticus. The blended mycelia from aerated cultures, grown for 8 to 10 days, have been shown to degrade AFB~ maximally, while 12-day-old mycelium from the cultures of A. parasiticus failed to degrade an appreciable amount of aflatoxin, suggesting thereby that some intracellular substance(s) is/are responsible for aflatoxin degradation/detoxification [40,44]. Harold and Smith [36] have observed that the intact mycelium of A. flavus could detoxify aflatoxins more efficiently than the cell-free extracts. However, the degradative ability of the extracts prepared from the older mycelium was significantly higher than the extracts of younger mycelium, suggesting that the levels of aflatoxin detoxifying enzymes appear to develop within the toxigenic fungus by an obscure mechanism.

Effect of p H and temperature on aflatoxin detoxification Biologically optimal pH and temperature have been quite instrumental in regulating the secondary metabolism in microorganisms. Doyle and Marth [37] observed that the 9000xg supernatant fraction of 9-day-old mycelium of A. parasiticus NRRL 2999 was able to detoxify aflatoxins B~ and G~ at pH 5.0 and 6.5. There was some aflatoxin degradation at pH 4.0, but essentially no degradation at pH 2.0 and 3.0. The fact, that such detoxffication at pH 5.0 and 6.5 was only biological, was confirmed by the finding that little or no chemical degradation of aflatoxins occurs at pH 5.0 and 6.5 [37,44]. The effect of temperature on detoxification of AFB~ and AFG~ was studied by Doyle and March [37] in A. parasiticus. They have shown that 9-day-old mycelia caused maximum aflatoxin degradation at 28~ with intermediate rates at 19~ and 36~ and little degradation at 45~ In contrast, Faraj [47] has shown that, when toxigenic A. flavus was grown at 30~ in either solid or liquid cultures, there was extensive aflatoxin synthesis, followed by the onset of ailatoxin degradation after approximately 5 days. Further incubation at 30~ gave continued aflatoxin detoxification. However, if cultures were transferred to high temperatures viz 40 ~ 45 ~ 50~ there were much increased rates of detoxification. Similar observations have been made by Singh and Smith [45] using the same toxigenic strain of A. flavus.

58 (ii) A f l a t o x i n d e t o x i f i c a t i o n b y t h e a t o x i g e n i c s t r a i n s of t h e same fungal species Doyle and March [41] and Marth and Doyle [44] have pioneered such studies by observing aflatoxin detoxification in A. parasiticus strains (toxigenic strain NRRL 2999 and atoxigenic strain NRRL 3315). They have compared the two strains and reported that the atoxigenic strain was less efficient in detoxifying the afiatoxins. An atoxigenic strain of A. flavus has been found to be most effective in detoxifying the aflatoxins produced by the toxigenic A. flavus and A. parasiticus strains under coculture conditions [ 4 8 ] . Nakazato et al. [48] have demonstrated that aflatoxin BR, produced by the toxigenic A. flavus and A. parasiticus, is transformed or metabolized by all strains of non-producing A. flavus; AF-A and AF-B were the common metabolites. Also, similar studies have already been carried out by Cotty [49] in cotton seeds. Not only that, the aflatoxin produced in A. flavus contaminated maize was degraded by the atoxigenic strain of A. flavus in solid-state fermentation [50]. This could be a very useful method of biological control of both preand p o s t - h a r v e s t aflatoxin c o n t a m i n a t i o n in a g r i c u l t u r a l produce, particularly the grains. (iii) Aflatoxin d e t o x i f i c a t i o n by o t h e r atoxigenic m i c r o o r g a n i s m s This approach to aflatoxin detoxification involves the use of microorganisms, other than toxigenic aspergilli, which do not themselves produce aflatoxins but apparently can metabolise them to less toxic or non-toxic molecules. Reduction of most hepatotoxic, hepatocarcinogenic and mutagenic afiatoxin (AFB 1) to a less toxic product, i.e., aflatoxin R 0 (aflatoxicol) has been observed in many organisms [51-53]. Biotransformation of AFB 1 to as yet uncharacterized compounds has also been reported with bacteria, including Corynebacterium rubrum and Lactobacillus spp., with the fungi - A . niger, Trichoderma viride,

Mucor ambiguos, M. alterans, Helminthosporium sativum, Rhizopus arrhizus, R. oryzae and R. stolonifer, and the p r o t o z o a n - Tetrahymena pyriformis [15,51,54-57]. However, the rate of conversion of AFB 1 to AFR O in Dactylium dendroides, Absidia repens, and Mucor grisseocyanus was very slow, taking 3-4 days to achieve only 60% reduction of AFB~ to AFR O [54]. Bol and Smith [58] studied detoxification of aflatoxin B~ by food grade Rhizopus strains. They observed that 87% of Rhizopus strains tested were positive to AFB 1 defluorescence on agar media. The isolates from c o n t a m i n a t e d feedstuffs showed d i m i n i s h e d (7.5%) defluorescence capacity. Out of 29 strains tested, 18 were able to eliminate 50-100% aflatoxin B 1 after 5 days incubation at 25~ Aflatoxin G 1 has also been found to be degraded by various Rhizopus spp. to an intermediate metabolite previously reported in A. flavus as aflatoxin B s and in A. parasiticus as parasiticol. Ciegler et al. [59] screened over 100 microorganisms for their ability to either degrade or

59 transform AFB 1 and found that Flavobacterium aurantiacum was most effective. F. aurantiacurn has further been shown to remove aflatoxins B1 and M 1 from liquid [52,53]. Knol et al. [60] have demonstrated that AFB~ could be eliminated from peanut meal by R. oryzae NRRL 395 in a solid-substrate process, which has been quite effective with major decreases (from 260 to 70 ~tg/kg) of AFB 1 content in this raw material. According to Shantha et al. [61], various strains of A. niger were able to decrease the levels of aflatoxin concentrations by about 64-99%, when they were grown with the toxigenic A. flavus in liquid cultures. The inhibition of AFB~-induced hepatocarcinogenesis by the Rhizopus delemer has been subject matter of the studies of Zhu et al. [12], and hence suggests the effective biodetoxification measure of aflatoxins. A. niger has also been shown to be most effective in reducing the levels of respective mycotoxins produced by A. ochraceus and A. flavus [62]. As well as, A. oryzae has been shown to reduce the production of aflatoxins in mixed cultures of this organism and the aflatoxigenic A. flavus [63]. Similarly, Choudhary [64] has demonstrated the inhibitory effects of Fusarium moniliforme, Trichoderrna viride, and Rhizopus nigricans on aflatoxin production, w h e n these microbes were co-cultivated with aflatoxin-producing fungus, A. flavus. The detoxification of aflatoxins in solid substrates is very important for both commercial and health reasons. Cuero et al. [65] have demonstrated that there was about 50% reduction in aflatoxin concentration, when A. flavus was grown with A. niger on maize samples. The decrease in total aflatoxin level was about 40%, 70%, and 75%, when this toxigenic fungus was cultured on maize with Fusarium graminiarum, A. oryzae and Penicillium viridicatum, respectively. Also, Cuero et al. [66] have implicated chitosan as well as the microbial agents, such as Bacillus subtilis and Trichoderma harzianum in biological control of aflatoxins in pre-harvest maize. Barrios-Gonzalez et al. [67] have shown, while e v a l u a t i n g the risk of aflatoxin c o n t a m i n a t i o n of cassava protein enrichment process with A. niger, that the toxigenic A. parasiticus can grow and produce aflatoxins under favourable environmental conditions such as suitable temperature, moisture content and nutrition. It was noticed that in mixed cultures, using A. niger and differrent amounts of A. parasiticus, the operation temperature of protein enrichment process (35~ drastically reduces the toxin production. Although nitrogen and phosphorus concentrations in the medium were partially inhibitory to aflatoxin biosynthesis, very high reduction could be attained. The best toxicological protection was by the atoxigenic strain itself (A. niger No.10). The aflatoxin production was completely inhibited when these two species (A. niger and A. parasiticus) grew together in solid-state fermentation, thereby suggesting t h a t the microorganisms, other than the toxigenic ones, can detoxify aflatoxins in the consumables (foods and feeds), and hence can present an effective measure of biological

60 control of aflatoxins, thus eliminating the risk to h u m a n health. CONCLUDING

and

animal

REMARKS

Aflatoxins are the serious source of contamination of most foods and feeds, thereby causing potential threat to both h u m a n and animal health. There has been a tremendous amount of information available on physical and chemical methods of detoxification of these potentially toxic, carcinogenic and mutagenic secondary metabolites. However, biodetoxification of aflatoxins by the factors already present in the substrates (such as the genetic constitution of the food commodity) infected with the toxigenic fungi, by the toxigenic as well as atoxigenic aspergilli or by various other microorganisms, provide a very useful, novel and safe method for biological control of these toxins. More attention should be given to the improved methods of decontamination and detoxification of the contaminated agricultural produce under natural solid-substrate conditions. AC K N O W L E D G E M E N T S

The help rendered by M s Deepika Mittal and Sandhya Singh as well as M r Rathendra Raman, Sudhir If~ Singh and Hemant I~ Singh is gratefully acknowledged. Thanks are also due to M r Lalit Kumar, M r Krishan Lal, M r Satish K u m a r Sundan, and M r S.K. Dass for technical help. REFERENCES

1 2 3 4 5 6 7

Smith JE, Moss MO. Mycotoxins : Formation, Analysis and Significance. Chichester, New York, Brisbane, Toronto, Singapore: John Wiley & Sons, 1985; pp.148. Cole RJ, Cox RH eds. Handbook of Toxic Fungal Metabolites. New York: Academic Press, 1981; pp.937. Borroz KI, Ramsdell HS, Eaton DL. Toxicol Lett (Amst.) 1991; 58(1): 97-106. Bryden WL, Cumm~ng RB, Lloyd AB. Avian Pathol 1980; 9: 539-550. CAST (Council for Agricultural Science and Technology) 1989; Task Force Report on Mycotoxins : Economic and Health Risk (No.l16). Iowa: CAST. Hynh VL, Gerdes RG, Lloyd AB. Aust J Biol Sci 1984; 37: 123-129. Keyl AC, Booth AM. J Am Oil Chem Soc 1971; 48: 599-604.

61 8 9 10 11 12 13

14 15 16 17

18 19 20 21 22 23 24 25 26 27 28 29 30 31

Lynch GP, Covey FT, Smith DF, Weinland BT. J Animal Sci 1972; 35: 65-68. Newberne PM, Butler WH. Cancer Res 1969; 29: 236-250. Newberne PM, Carlton WW, Wogan GN. Pathol Vet 1964; 1: 105132. Tung TC, Ling KH. J Vitaminol 1968; 14: 48-52. Zhu C, Min-Jie D, Dao-Nian L, Lue-Queen W. Mater Med Pol 1989; 21(2): 87-91. Bilgrami KS, Sinha KK. In : Mukerji KG, Pathak NC, Singh VP, eds. Frontiers in Applied Microbiology. Lucknow: Print House India, 1985; 1: 349-361. Kurtzman CP, Horn BW, Hesseltine CW. Antonie van Leeuwenhoek 1987; 53: 147-158. Singh VP, Mukerji KG. In 9 Mukerji KG, Singh VP, eds. Concepts in Applied Mirobiology and Biotechnology. New Delhi: Aditya Books Pvt Ltd, 1994; (in press). Wong JJ, Hsieh DPH. Proc Natl Acad Sci USA 1976; 73: 2241-2244. IARC (International Agency for Research on Cancer). IARC Monographs on the Evaluation of Carcinogenic Risk of Chemicals to Man. Lyon: International Agency for Research on Cancer, 1976; 10: 51-72. Purchase IFH ed. 1974. Mycotoxins. Amsterdam: Elsevier. Goldblatt LA, DoUear FG. Pure Appl Chem 1977; 49: 1759-1764. Shrivastava AK, Ranjan KS, Ansari AA. J Food Sci Technol 1991; 28(3): 189-190. Samarajeewa U, Sen AC, Fernando SV, Ahmed EH, Wei CI. Food Chem Toxicol 1991; 29(1): 41-48. Jorgensen KV, Park DL, Rua SN Jr, Price RL. J Food prot 1990; 53(9): 777-778. Abdel-Rahim EA, Naguib KM, Badawi MM, Ibrahim MKK, Guergues SN. Grasas Aceites 1990; 41(2): 144-148. Mahalingam RJ, Govindan S, Punniamurthy N, Balachandran C. Indian Vet J 1990; 67(2): 149-151. Altug T, Yousef AE, Marth EH. J Food Prot 1990; 53(7): 581-582. Howes AW, Dutton MF, Chuturgoon AA. Mycopathol 1991; 113(1): 25-29. Zeringue HJ Jr, Bhatnagar D. J Am Oil Chem Soc 1990; 67(4): 215216. Costa JL, Da S, Kushalappa AC. Summa Phytopathol 1989; 15(2): 156-162. Azaizeh HA, Pettit RE, Saar BA, Phillips TD. Mycopathol 1990; 110(3): 125-132. WaUin JR, Windstrom NW, Fortnum BA. J Sci Food Agric 1991; 54(2): 235-238. Song D. Acta Microbiol Sin 1991; 31(3): 169-175.

52 32 33 34. 35 36 37 38 39

40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57

Rasic JL, Skrinjar M, Markov S. Mycopathol 1991; 113(2): 117119. Patterson DSP. Food Cosmet Toxicol 1973; 11: 287-294. Raney KD, Shimada T, Kim D, Groopman JD, Harris TM, Guengerich FP. Chem Res Toxicol 1992; 5(2): 202-210. Daniels JM, Lui L, Stewart RK, Massey TE. Carcinogenesis (Lond.) 1990; 11(5); 823-828. Harold AB, Smith JE. J Gen Microbiol 1987; 113: 2023-2029. Doyle MP, Marth EH. Eur J Appl Microbiol Biotechnol 1978; 6: 95100. I~peli O. Microbiol Rev 1986; 50: 244-258. Ruckpaul K, Rein H, Blanck J. In : Ruckpaul K, Rein H, eds. Frontiers in Biotransformations (Vohl) : Basic Mechanisms of Regulation of Cytochrome P-450. London, New York, Philadelphia: Taylor & Francis, 1989; 1-65. Doyle MP, Marth EH. J Food Prot 1978; 41: 549-555. Doyle MP, Marth EH. Mycopathol 1978; 63: 145-153. Doyle MP, Marth EH. Mycopathol 1978; 64: 59-62. Doyle MP, Marth EH. Eur J Appl Microbiol Biotechnol 1979; 7: 211-217. Marth EH, Doyle MP. Food Technol 1979; 33: 81-87. Singh VP, Smith JE. Biotechnological implications of high temperature metabolism in microorganisms. In: Proc Summer Conf Soc Appl Bacteriol, Bristol (UK), 1991; 50. Shih CN, Marth EH. Z Lebensm Unters-Forsch 1975; 158; 361-362. Faraj MK. Regulation of Mycotoxin Formation in Zea mays. Ph.D. Thesis, University of Strathclyde, Glasgow (UK), 1990. Nakazato M, Morozumi S, Saito K, Fujinuma K, Nishima T, Kasai N. Risei Kagaku 1991; 37(2): 107-116. Cotty PJ. Plant Dis 1990; 74(3): 233-235. Brown RL, Cotty PJ, Cleveland TE. J Food Prot 1991; 54(8): 623626. Cole RJ, Kirksey JW, Moore JH, Blankenship BP, Diener UL, Davis ND. Appl Microbiol 1972; 24: 248-256. LiUehoj EB, Ciegler A, Hall HH. Can J Microbiol 1967; 13: 624627. Lillehoj EB, Stubblefield RD, Sham_one GM, ShotweU OL. Mycopathol Mycol Appl 1971; 45: 259-264. Detroy RW, Hesseltine CW. Nature 1968; 219: 967. Detroy RW, Hesseltine CW. Can J Biochem 1970; 48: 830-832. Mann R, Rehm HJ. Eur J Appl Microbiol Biotechnol 1976; 2: 297306. Robertson JA, Teunisson DJ, Boudreaux GJ. J Agric Food Chem 1970; 18: 1090-1091.

53 58 59 60

61 62 63 64 65 66 67

Bol J, Smith JE. Food Biotechnol 1989; 3: 127-144. Ciegler A, LiUehoj EB, Peterson RE, Hall HH. Appl Microbiol 1966; 14: 934-939. Knol W, Bol J, Huis In T, Yeld JHJ In : Zeuthen P, Chei~el JC, Erikson C, Gormley TR, Liko P, Paulus K, eds. Processing and Quality of Food. London, New York: Elsevier Applied Science, 1990; 2: 2.133-2.136. Shantha T, Rati ER, Bhawani Shankar TN. Antonie van Leeuwenhoek 1990; 58(2): 121-128. Paster N, Pushinsky A, Menasherov M, Chet I. J Sci Food Agric 1992; 58(4): 584-592. Sardjono RK, Sudarmadji S. Asian Food J 1992; 7(1): 30-33. Choudhary AI~ Lett Appl Microbiol 1992; 14(4): 143-147. Cuero R, Smith JE, Lacey J. J Food Prot 1988; 51: 452-456. Cuero RG, Duffus E, Osuji G, Pettit R. J Agric Sci 1991; 117(2): 165-170. Barrios-Gonzalez J, Rodriguez GM, Tomacini A. J Ferment Bioeng 1990; 70(5): 329-333.

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Biotransformations: Microbial Degradation of Health Risk Compounds Ved Pal Singh, editor 9 1995 Elsevier Science B.V. All rights reserved.

Metabolism and C-2 h y d r o c a r b o n s

cometabolism

of h a l o g e n a t e d

65

C-1 a n d

Mukesh I~ Jain a and Craig S. Criddle b ~ Michigan

of Civil and Environmental Engineering, State University, East Lansing, MI 48824, U.S.A.

bNational Science Foundation Center for Microbial Ecology, Michigan State University, East Lansing, MI 48824, U.S.A. INTRODUCTION Chlorinated hydrocarbons, containing one or two carbon atoms, constitute a significant fraction of the h a z a r d o u s substances from industrial, domestic, and agricultural sources. In part, this is due to their high levels of production. Over five million tonnes of 1,2dichloroethylene (1,2-DCE) are produced annually for use as a solvent and chemical intermediate [1]. Vinyl chloride (VC) is also produced in large amounts (over three million tonnes annually) for the manufacture of polyvinyl chloride [1]. The solvents tetrachloroethylene (PCE), trichloroethylene (TCE), 1,1,1-trichloroethane (TCA), 1,1dichloroethylene (1,1-DCE), 1,2-dichloroethane (1,2-DCA), and carbon tetrachloride (CT) have a combined annual production of over 6 million tonnes [1]. Since 1970, annual U.S. production of dichloromethane (DCM) has ranged from 212 to 286 million kg, with the principal application being paint removal [2]. One- and two- carbon halogenated compounds tend to be mobile and persistent in soils and ground waters [3]. Among the most commonly detected ground water contaminants are PCE, TCE, TCA, 1,1-DCE, 1,2DCA, CT, chloroform (CF) and certain chlorofluorocarbons (CFCs). Many of these chemicals are classified as priority pollutants by the United States E n v i r o n m e n t a l Protection Agency (USEPA), and are known or suspected carcinogens or mutagens. Some have potential for ozone depletion. Release to the environment is caused by inadequate disposal techniques, accidents, deliberate agricultural applications, or chlorination of water and wastewater. Removal of C-1 and C-2 chlorinated aliphatics from w a t e r by physicochemical processes such as carbon adsorption or air stripping transfers contaminants from the aqueous phase to a solid or gaseous phase. In contrast, biological processes can destroy contaminants. To be effective, however, conditions t h a t favour growth of a transforming population must be created (biostimulation) or the transforming organisms must be added (bioaugmentation). In addition, care must be taken to

66 p r e v e n t or m i n i m i z e t r a n s f o r m a t i o n s t h a t yield i n t e r m e d i a t e s or byproducts that are hazardous. In this article, we summarize research on C-1 and C-2 haloaliphatic compounds with a focus on the agents of transformation (in the absence of light), growth kinetics, transformation kinetics, and pathways of transformation.

R E A C T I O N TYPES Reactions affecting the environmental fate of halogenated one- and two-carbon compounds can be broadly classified as substitutions, dehydrohalogenations, oxidations, and reductions [4]. These reactions can be either abiotic or biotic. Dehydrohalogenations are typically abiotic, while oxidations in dark e n v i r o n m e n t s are mostly biotic. Substitutions and reductions can be either biotic or abiotic. With some notable exceptions, abiotic transformations tend to be slow. Biotic transformations can be rapid when the microorganisms, that synthesize reactive enzymes or cofactors, are present in sufficient numbers. Several of the less halogenated aliphatic compounds (dichloromethane, 1,2-dichloroethane, etc.) are good electron donors and can serve as growth substrates. Usually, these compounds are susceptible to initial attack by oxidation or hydrolysis. The products of these reactions are typically alcohols or acids, that can be further oxidized by the transforming population to give carbon and energy for growth and maintenance. There is also some evidence that certain highly halogenated aliphatic compounds (such as tetrachloroethylene) can serve as the terminal electron acceptors for growth for some organisms [5]. Usually, though, the highly halogenated aliphatics do not support growth and are transformed only by cometabolism. Cometabolism is defined here as the transformation of a nongrowth substrate by growing cells in the presence of growth substrate, by resting cells in the absence of growth substrate, or by resting cells in the presence of energy substrate [6]. A growth substrate is defined as an electron donor, that supports growth. An energy substrate is defined here as an electron donor, that provides reducing power and energy for the transforming population, but does not, by itself, support growth. Cometabolism results from the lack of specificity of enzymes and cofactors. The products of cometabolic reactions accumulate in pure cultures, but, in a mixed culture, they are typically used by other microorganisms. As a result, cometabolic transformations are key initiatory reactions in pathways, that ultimately result in the complete degradation of many hazardous chemicals [7]. The first known examples of cometabolism were all oxidations, and, as a result, the t e r m "co-oxidation" was used to describe them. Subsequently, reductive transformations were discovered, that did not facilitate growth of the transforming orga_nisms, and depend upon the

67 concurrent or previous utilization of a growth or energy substrate. These "co-reductions" led to the use of the broader term, cometabolism. It now appears that certain cometabolic reactions are also hydrolytic. Thus, in addition to the well-known examples of co-oxidation, we now recognize the potential for "co-reductions" and "co-hydrolyses". All of the known co-oxidations occur only under obligate aerobic conditions, while most co-reductions occur under anaerobic conditions.

Substitution The type reaction

for substitutions is: RX + N - - - - > RNu + X~

(1)

In the above reaction, RX is an alkyl halide and Nu is a nucleophile. The most important nucleophile is water. Reactions with water result in replacement of a halogen by -OH (hydrolysis). Halogenated a|iphatic compounds undergo hydrolysis in the absence of inorganic or biochemical catalysts. Abiotic hydrolysis reactions are bimolecular, with water as the d o m i n a n t nucleophile, but because w a t e r is p r e s e n t at high concentrations, pseudo-first-order kinetics are observed. Many hydrolysis reactions are potentially faster at higher pH, where the hydroxide ion acts as the nucleophile. However, below pH 11, a pH dependence for substitution reactions is generally not observed [8,9]. The nature of the h a l o g e n s u b s t i t u e n t s and t h e degree of h a l o g e n a t i o n i n f l u e n c e s substitution rate. Increased halogenation leads to slower substitution reactions and longer half lives [8,9]. In general, abiotic substitution reactions proceed slowly, but can be g r e a t l y a c c e l e r a t e d by enzymes. E n z y m e - m e d i a t e d substitutions frequently involve cysteine residues in proteins or peptides, such as glutathione. Biotic and abiotic hydrolysis of h a l o g e n a t e d aliphatic compounds yields alcohols by hydroxyl substitution at the halogenated carbon [10]. If t h e s e alcohols are themselves halogenated, f u r t h e r hydrolysis to acids or diols can occur. Examples of microbially-mediated hydrolysis reactions, together with responsible enzymes, are provided in Table 1.

Dehydrohalogenations The

type

reaction I I --C--C-, , H X

for

-

dehydrohalogenation

---

\ /

C=C

/ \

+

HX

is:

(2)

68 Table 1 Examples of hydrolysis and co-hydrolysis Microorganism

Hydrolysis-

Growth substrate/ nongrowth substrate

Responsible enzyme(s)

Refs

dichloromethane

halidohydrolase

[ 104 ]

dichloromethane dibromomethane

glutathione+ glutathione-Stransferase

[97]

1,2-dichloroethane

haloalkane dehalogenase and haloacid dehalogenase

[75,76]

dichloromethane dibromomethane

DCM dehalogenase group A

[105]

1-haloalkanes

dehalogenase

[106]

dichloromethane dibromomethane

DCM dehalogenase group B

[98]

amino acids/ 1,1,1trichloroethane

unknown enzymes

[71]

acetate, glycerol/ carbon tetrachloride

unknow~ iron scavenging agent

[82,83]

metabolism

Pseudomonas DM1 and DM2

Hyphomicrobium DM2

Xanthobacter autotrophicus GJ10 Ancylobacter aquaticus AD20 and AD25

Methylotrophic bacterium sp. strain DM4

Arthrobacter sp. strain

HA1

Methylotrophic bacterium strain DMll

[74]

Co-hydrolysis Clostridium TCAIIB

Pseudomonas sp. strain KC

For halogenated aliphatics, dehydrohalogenations are abiotic. Polychlorinated alkanes undergo dehydrohalogenation under extreme basic conditions, and at p H 7 [11]. These reactions generally follow bimolecular kinetics, depending on hydroxide ion concentration. At

69 n e u t r a l pH conditions, dehydrohalogenation by weaker bases (e.g., water) might be important. The number and kind of halogen substituents have a strong influence on d e h y d r o h a l o g e n a t i o n rates. I n c r e a s e d halogenation tends to decrease substitution reaction rates (hydrolysis) and increase dehydrohalogenation rates. Consequently, highly halogenated C-2 alkanes are susceptible to dehydrohalogenation, except, of course, those that are fully halogenated. A few compounds undergo simultaneous dehydrohalogenation and hydrolysis [12]. A well-known example is 1,1,1-TCA, which undergoes simultaneous hydrolysis to acetate and dehydrohalogenation to 1,1-dichloroethylene [13]. Oxidations

Type reactions (a) (b)

(c)

for aerobic

monooxygenase I

I

I

0 2 + 2 H + + 2e- + / C : C

+

2H +

~

are:

I

02 + 2H + + 2e" + -C-H

:2.

oxidations

+ 2e- + ,C=C

"*

/ "

'

~

monooxygenase

dioxy_genase

.....

=

-'C-OH

+ H20

(3)

O

~ C-C , HO '

+H

2,

O

OH '

-C-CI

9

I

Aerobic oxidations rely upon the catalytic activity of nonspecific monooxygenases or dioxygenases. As indicated in the type reaction sequences, these enzymes require both reducing power and molecular oxygen. They are widely distributed in n a t u r e in many microbial populations, including methanotrophs, nitrifiers, numerous hydrocarbon degraders, and they are even found in higher organisms, including man. Frequently, oxygenase reaction products are not useful to the transforming organisms, so many oxidative transformations are cometabolic. Several examples of co~xidizing bacteria are provided in Table 2. Oxygenases catalyze the incorporation of oxygen, derived from molecular oxygen, into the halogenated molecule. As shown in reaction 3a, oxygen may be inserted into the carbon-hydrogen bond creating halogenated alcohols, that spontaneously eliminate HX to give an aldehyde [14,15]. As shown in reaction 3b, oxygen may also be inserted into carboncarbon double bonds yielding an epoxide [14]. Halogenated aldehydes or acyl chlorides are common intermediates, and are typically oxidized or hydrolyzed to acids or reduced to alcohols [16-18]. Halogenated molecules, t h a t are oxidized include hydrogen-containing alkanes and alkenes. Completely halogenated alkanes and alkenes are resistant to oxidation by oxygenases.

70 Table 2 Examples of co-oxidation of alkyl halides Microorganisms

Growth substrate

Pseudomonas cepacia strain G4

phenol, toluene T C E and o-cresol phenol

[53,54]

Pseudomonas putida F1

toluene

TCE

[55,56]

Strain

toluene

TCE

[22,59]

methane methanol formate

TCE chloroalkanes except CT chloroalkenes except PCE

[20,109] [23] [62]

methane

TCE chloroalkenes except PCE chloroalkanes except CT

[64] [63,110]

Mycobacterium vaccae JOB5

propane

chloroalkenes PCE

[111 ]

Nitrosomonas europaea

ammonia

TCE chloroalkenes except PCE chloroalkanes except CT

[50,51, 112]

Xanthobacter strain

propylene

TCE

[ 113,114]

toluene

TCE

[114]

46-1

Meth ylosinus trichosporium OB3b Methylocystis strain M

sp.

Nongrowth substrate

Refs

[107,108]

except

Py2 Genetically engineered Escherichia coli

Nonspecific oxygenases that figure prominently in the process of cooxidation have great potential for degradation of halogenated aliphatic compounds. Among the most important are the methane monooxygenases (MMOs) found in methanotrophs. These enzymes endow the methanotrophs with the ability to oxidize virtually all of the halogenated aliphatic

71 hydrocarbons, with the exception of those that are completely halogenated. All methanotrophs tested are able to form a particulate type MMO (pMMO) or membrane-bound enzyme, whereas some cultures grown under copper limitation are capable of producing a soluble type of MMO (sMMO), with a broader substrate range than pMMO [19,20]. When induced for sMMO, Methylosinus trichosporium OB3b co-oxidized all chlorinated aliphatic hydrocarbons (C1 to C3) except PCE and CT [2023]. For methanotrophs, formate can serve as an energy substrate, increasing both the rate and extent of TCE cometabolism. Although methanotrophs obtain reducing power from the oxidation of formate to carbon dioxide, they are unable to assimilate formate, and they can not use it as a growth substrate.

Reductions The type reactions for reductions are:

(a)

(b)

(c)

I

-C-X

I

+ X"

I

(4)

+ H+§ e- -..-.- - C - H

I

(hydrogenolysis)

1

I

I

1

*

--C-C--

\Cffic'X+ /

I

I

-C.

X

(d)

+e- ---.--C-

\

+r

=

H ++ 2r -

f\ C f C ~

/

+ X"

/ j \C _ - - C %

(di-halo

elimination)

+ X"

Table 3 lists pure cultures, capable of co-reducing halogenated aliphatic compounds. Frequently, t h e s e o r g a n i s m s possess t r a n s i t i o n metal complexes that react with the alkyl halides. Among the most important of these complexes are the cytochromes, corrinoids, factor F430, and vitamin B m [24]. Reductions by transition metal complexes are typically initiated by the transfer of a single electron, loss of a single halide substituent, and the formation of a free radical (reaction 4a). Formation of the free radical is the first and, in most cases, the rate-limiting step in the reduction of halogenated aliphatic compounds. The free radical can undergo a range of reactions, depending upon the nature of the radical and its environment. If the radical abstracts hydrogen from water or

72 Table 3 Examples of co-reduction of alkyl halides Microorganism

Growth substrate

Nongrowth substrates

Refs

amino acids

1,1,1-trichloroethane carbon tetrachloride chloroform

[71]

DCB-I

chlorobenzoate

tetrachloroethane

[38]

Methanobacterium thermoautotrophicum

acetate, methanol

1,2-dichloroethane

[37,77]

Methanosarcina Methanosarcina strain DCM

mazei sp.

acetate, methanol

tetr achloroethane trichloroethane

[36,39 ]

Methanosarcina strain DCM

sp.

methanol

chloroform

[92 ]

Methanosarcina barkeri

H2-CO 2

I,2-dichloroethane chloroethane

[78 ]

Desulfobacterium autotrophicum

lactate

carbon tetrachloride I,1,l-trichloroethane

[37]

Acetobacterium

fructose

carbon

tetrachloride

[81]

Escherichia coli k12 (fermenting)

glucose

carbon

tetrachloride

[79]

Escherichia coli k12 (fumarate-respiring)

glycerol

carbon

tetrachloride

[79]

Clostridium

sp. TCAIIB

woodii

from a surrounding organic, the product is similar to the parent compound, with a halogen replacing one of the hydrogen substituents (hydrogenolysisreaction 4b). If the carbon radical is located adjacent to a halogenated carbon, then a second halide can be lost with formation of a double bond (di-halo elimination - reaction 4c). In general, the greater the degree of halogen substitution, the more oxidized a molecule becomes, and the more susceptible it is to reduction by biotic or abiotic electron donors [4]. Alkyl halide mixtures are susceptible to reductive transformation under anaerobic conditions [3,4,25-28]. In particular, Bouwer and McCarty [3,29] and Vogel et al. [4] reported that mixtures of 1,1,1-TCA; 1,2-DCA;

73

TCE; 1,2 dibromoethane, and P C E are transformed in methanogenic environments. Anaerobic habitats, supporting reductive transformation, include continuous flow methanogenic fixed-film reactors [3,4,30,31], organic sediment-muck [24,27,32-34], anaerobic aquifer microcosms [28], anaerobic sediment from the Rhine river [35], enrichments [26], and pure cultures [36-39].

KINETICS OF GROWTH If the concentrations of all but one of the substances needed for bacterial growth are present at levels, that exceed growth requirements, then the limiting substrate is termed as the growth-limiting substrate, or simply, the growth substrate. For growth substrates, the rate of substrate utilization is a function of the growth rate of the microorganisms. A widely used model of bacterial growth and decay is the Monod expression as modified by Herbert et al. [40]: dX/dt ~t =

YkS ffi Y q g - b =

X

-b

(5)

K+S

where: ~t ffi specific growth rate, da~r 1 X ffi o r g a n i s m concentration, rag/1 t = time, days y = maximum organism yield, mg cell/rag substrate q~ ffi specific rate of utilization of growth substrate, mg growth s u b s t r a t e / m g cell-d b = decay coefficient, day -1 k = maximum specific rate of substrate utilization, mg growth s u b s t r a t e / m g cell-d S = growth substrate concentration, rag/1 K ffi half saturation coefficient for the growth substrate, mg growth substrate/1 W h e n S is zero, concentration decreases. of growth substrate at is found from equation

Smio=

m i c r o o r g a n i s m s u n d e r g o decay, and b i o m a s s A concentration of interest is the concentration which growth and decay are equal ( S n ) , which 1 by letting tt = 0 [41]:

K

Yk-b

(6)

74 When the concentration of a growth substrate is above Smin, net growth will occur. Under such conditions, the substrate is termed a primary substrate. When the concentration is below Stain, then the substrate is termed a secondary substrate. If a primary substrate is available, or if the collective concentration of secondary substrates permits growth, then the concentration of a secondary substrate can be reduced to a value significantly below its S n . Under such conditions, even low concentrations of a substrate, like those found in many ground waters, can be removed [7]. Values of k, K , and hence k' for secondary substrates may differ from those of the primary substrate. In some cases, k' of the secondary substrate is greater than k' of the primary substrate, and in other cases it is much less. When the k' values for the secondary substrate are much less t h a n k' values for the primary substrate, then large amounts of primary substrate may be required to degrade the secondary substrate.

KINETICS

OF T R A N S F O R M A T I O N

Substrate transformation kinetics Transformations of growth substrate by described using saturation kinetics [42]"

-dS/dT q ffi

bacteria

can

generally

be

kS ffi

X

(7) K+S

Transformations of nongrowth substrate by resting cells can also be described using saturation kinetics: qc=

-dC/dT X

=

k C K+C

(8)

where" qr = specific rate of transformation of nongrowth substrate, mg s u b s t r a t e / m g cell-d k ffi m a x i m u m specific rate of nongrowth substrate utilization, mg substrate/mg cell-d C = concentration of nongrowth substrate, rag/1 K = half saturation coefficient for the nongrowth substrate, mg substrate/1. If the substrate concentration is low (S